Paracetamol (N-acetyl-para-aminophenol) disrupts early human embryogenesis

doi:10.21203/rs.3.rs-3685511/v1

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Paracetamol (N-acetyl-para-aminophenol) disrupts early human embryogenesis

https://doi.org/10.21203/rs.3.rs-3685511/v1

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It is estimated that 10–40% of all human conceptions fail around implantation^1-7. Genetics explain ≈ 50% of early embryonic loss, leaving a substantial part of early loss without a known cause^8,9. Smoking and alcohol are known risk factors for spontaneous abortion, indicating the importance of the chemical environment during embryonic development¹⁰. Here we show that paracetamol (N-acetyl-para-aminophenol (APAP); otherwise known as acetaminophen), the recommended medication for pregnant people for treatment of mild to moderate pain and fever¹¹ and an environmental pollutant^12-15, disrupts both mouse and human pre-implantation development. We found that APAP inhibited cell cycle progression, likely through ribonucleotide reductase, resulted in blockage of DNA synthesis across all model systems, and reduced pregnancy outcomes in mouse models. At concentrations found in the reproductive system of women after standard administration, APAP exposure decreased human cleavage stage embryo cell numbers or caused direct embryonic fatality. Similar exposure to human blastocyst stage embryos for 6 h resulted in decreased DNA synthesis as well as morphological changes. Our data demonstrate that a widely used mild analgesic and environmental pollutant might result in embryonic loss and provide a foundation for understanding environmentally caused cell cycle inhibition in other processes during development.

Biological sciences/Developmental biology/Cell proliferation

Biological sciences/Developmental biology/Embryogenesis

Biological sciences/Molecular biology/Cell division/DNA replication/DNA synthesis

Health sciences/Diseases/Reproductive disorders/Infertility

Paracetamol/acetaminophen

cell cycle

early embryo development

pregnancy loss

A considerable percentage of human conceptions are lost before birth^1,2. Consistent data from the last decades have shown 10-40% of all embryos fails to implant and further 10-15% pregnancies are lost after implantation before clinical recognition^1–7,16,17. Studies using pre-implantation genetics have shown that chromosomal abnormalities are likely responsible for ≈50% of peri-implantation loss^8,9, suggesting that environmental factors might play a role in the loss of pregnancies^10,17. Smoking and alcohol are already known risk factors for spontaneous abortion, indicating the importance of the chemical environment during early development¹⁰. Recent studies also indicate that the vaginal microbiota play a role during development, especially among certain ethnic groups^18–21. While these studies have yielded novel insights into preterm labour, they do not fully explain the risk of early embryonic loss within the first weeks of pregnancy.

Over-the-counter mild analgesic paracetamol (N-acetyl-para-aminophenol (APAP), otherwise known as acetaminophen) is the active pharmaceutical ingredient in more than 600 different medications used to relieve mild to moderate pain and reduce fever¹¹. APAP is widely used by women of reproductive age and during pregnancy, as the compound has long been considered to be of minimal risk for use during pregnancy, when used as directed^22–24. Although studies have shown that APAP exposure at non-cytotoxic levels inhibits cell proliferation, impairs DNA repair, and increases DNA fragmentation^25–28, up to 65% of pregnant women use APAP in the USA, while the worldwide estimate is 50%^29–32. Moreover, APAP is a frequent pollutant of the world’s rivers and waterways¹⁵ and ubiquitous environmental background exposure to APAP has been found in several European countries^12–14.

We investigated whether APAP interferes with cell division and pre-implantation development (PID). To accommodate the challenges of determining the mechanism of action and effect of APAP during PID, we employed a variety of in vitro, ex vivo, and in vivo approaches in model organisms and humans ranging from yeasts to the human embryos and women at fertile age. Our results show that APAP restricts cell proliferation via inhibition of DNA synthesis and that exposure during the PID can lead to embryonic loss.

APAP inhibits cell proliferation, S-phase progression, and DNA synthesis in human somatic and embryonic stem cell lines

We exposed human embryonic kidney (HEK293) cells to 500 µM APAP. This resulted in decreased total cell numbers in a time-dependent manner without effecting cell viability (Supplemental Fig. 1a-b, n=6). The cells exposed to APAP displayed accumulation in the S phase of the cell cycle with a significant fold change of ≈1.5 as compared to the control (Supplemental Fig. 1c, n=6), concomitantly with reduced de novo DNA synthesis (Supplemental Fig. 1d, n=5-6).

Human embryonic stem cells (hESCs) are pluripotent cells derived from the inner cell mass (ICM) of a pre-implantation blastocyst stage embryo and a validated model system of PID³³. Exposure of hESC lines H1 and HUES4 to 200 µM APAP resulted in similar effects as observed in the somatic HEK293 cells (Supplemental Fig. 1e-f, i-j, n=5), including a time-dependent reduction in the total cell number without effecting viability, S phase accumulation (Supplemental Fig. 1g, k, n=5) and reduction of de novo DNA synthesis as compared to controls (Supplemental Fig. 1h, l, n=4). In addition, we found a reduction of cells in the G2/M phase, reflecting accumulation in the S-phase. Taken together, these data show that APAP restricts cell proliferation via inhibition of DNA synthesis.

APAP inhibits ribonucleotide reductase in yeast

APAP has been reported to cause DNA replication stress presumably through inhibition of ribonucleotide reductase (RNR)²⁷, an essential enzyme for DNA synthesis that reduces ribonucleotides to 2’-deoxyribonucleotides, the building blocks of DNA. As RNR is highly conserved across the eukaryotic kingdom³⁴, we tested the effect of APAP on RNR in three strains of fission yeast Schizosaccharomyces pombe (S. pombe), a model organism that does not express pharmaceutical APAP targets as prostaglandin-endoperoxide synthases (PTGS1 and PTGS2) and cannabinoid receptor 1 (CB1)³⁵. The first strain was deleted for the ddb1-gene (ddb1∆), which results in reduced 2’-deoxyribonucleotide pool and increased sensitivity to chemical RNR inhibition as compared to wild type (WT) cells. The second strain was deleted for both the ddb1-gene and the RNR inhibitory gene spd1 (ddb1Δ spd1Δ), which results in increased RNR activity and thus a ≈2-fold increased 2’-deoxyribonucleotide pool as compared to WT cells. The third strain was deleted for the ddb1-gene together with an activating point mutation in the large RNR subunit encoding gene cdc22 (ddb1Δ cdc22-D57N), resulting in a ≈5-fold increased 2’-deoxyribonucleotide pool as compared to WT cells³⁶. The RNR restricted ddb1∆ strain had decreased proliferation at 40- and 60 mM APAP as compared to WT. In contrast, this effect on cell division was suppressed in both the ddb1Δ spd1Δ and ddb1Δ cdc22-D57N strains with increased RNR activity (Supplemental Fig. 2a).

To understand the effect of APAP on human RNR, we used a recombinant S. pombe strain with the native RNR coding regions cdc22 and suc22 replaced with cDNA encoding the human orthologous RRM1 and RRM2 subunits (hR1/R2). Compared to the WT strain, this strain was more sensitive to APAP in terms of growth, both on solid agar and in liquid culture (Supplemental Fig. 2b-c; (c, n=3)), correlating with accumulation of cells in G1- and S-phase (Supplemental Fig. 2d). Taken together, these data are consistent with an inhibitory action of APAP on RNR, resulting in disruption of cell cycle progression at the G1/S-border and through the S-phase.

APAP delays early preimplantation development in mice

To understand the effect of APAP on PID, we investigated the effect of 24 or 48 h exposure (10-200 μM APAP) using C57BL/6 mouse 2 cell embryos (embryonic day (E)1.5). After 24 h of development, controls (n=21) as well as embryos exposed to 10 µM (n=12) consisted of the expected 8 blastomeres, whereas embryos exposed to 25 μM (n=11) had an equal distribution of 4 or 8 blastomeres. All embryos exposed to 50-200 μM APAP (n=6-9) consisted of 2-4 blastomeres, suggesting a delay in the 2^nd to 3^rd cleavage stages (Fig. 1a, d). After 48 h, most control embryos had developed into early or expanded blastocysts that were identified by a well-defined fluid-filled cavity (blastocoel). No embryos exposed to 50-200 μM (n=7-11) developed beyond the early blastocyst stage and all had reduced cell number as compared to controls (n=15) (Fig. 1a, e and Supplemental Fig. 3a). Blastomere morphology was not affected after 24 h of APAP exposure compared to control except for embryos exposed to 200 μM of APAP (n=8); here a minority of embryos were amorphic with fragmented blastomeres (Fig. 1b).

Among embryos in the control group cultured for 48 h, we observed more OCT3/4-positive cells (marker for ICM) relative to CDX2-positive cells (marker for trophectoderm, TE) with some cells identified with both markers as expected for this developmental stage of lineage specification³⁷. Among the low APAP exposure groups (10-25 µM, n=11-12), we observed a similar differentiation of the ICM and TE with cells expressing both markers of ICM and TE. In contrast, APAP concentrations >25 µM displayed fewer blastomeres with exclusive TE (CDX2 positive) cells as compared to the control embryos (Fig. 1c). Moreover, APAP-exposed embryos displayed cell fragmentation and amorphic nuclei with increasing severity and frequency at the higher concentrations of 150 and 200 μM (Fig. 1c). Calculating a growth rate based on the 48 h experiments, we found that APAP delayed development, increasing cellular doubling time from ≈13 h for controls to ≈30 h for 200 µM APAP exposed embryos (Fig. 1f). Moreover, delayed embryonic development was also reflected in a decreased blastocyst rate (Supplemental Fig. 3a). Together these experiments demonstrate that APAP delays or disrupts embryonic development in the mouse in a time and concentration dependent manner.

APAP exposure reduces cell number of the inner cell mass of mouse blastocyst

To understand the effect of APAP during later stages of PID, we exposed early mouse blastocysts (E3.5) to 100 (n=14), 150 (n=21) and 200 µM (n=26) APAP for 24 h. Most APAP-exposed embryos continued blastocyst development with a clearly defined expanded blastocoel, TE, and ICM (Fig. 2a) and with cell numbers equivalent to those of controls (n=26) (Fig. 2b-c). However, the number of cells in the ICM was decreased after 200 μM APAP (n=19) as compared to controls (n=15) (Fig. 2b, d); a phenotype that was recapitulated in embryos stained for microtubules and F-actin, displaying reduced ICM size while the overall structural organization of cells was maintained (Fig. 2e). Taken together, these data suggest that APAP delays the expansion of the pool of pluripotent ICM cells during the final stage of PID (E3.5-E4-5).

APAP during early mouse preimplantation development reduces number of full-term fetuses

To evaluate the effect of APAP exposure on implantation rate and subsequent development, we performed embryo transfer to pseudo-pregnant recipient dams with exposed and control embryos placed in each of the uterine horns, respectively (Fig. 3a). Exposing 2-cell embryos (E1.5) for 24 h to 100 or 200 µM APAP (n=4-5) did not affect the implantation rate, but decreased number of full-term fetuses and increased resorption points at 200 µM as compared to controls (Fig. 3b-c). Increasing the exposure time to 48 h decreased implantation rate and decreased the number of resorption points and full-term fetuses in a concentration-dependent manner with an impact on all three experimental endpoints (implantation, full term fetuses, and resorption points) as compared to controls (Fig. 3d-e, n=3). These results suggest that disrupted PID after exposure to APAP impact the ability of embryos to implant and/or survive after implantation.

Intrauterine APAP exposure results in an increase of resorption points and a decrease in full-term fetuses in mice

To understand the effect of intrauterine exposure on PID and subsequent development, we gavaged C57/Bl6 dams with APAP (200 mg/kg/day) from 1 day post coitum (dpc) and the subsequent 10 days (Fig. 4a, n=19-20). At caesarean section at 18 dpc, dams treated with APAP had reduced number of live fetuses. Moreover, post-implantation embryonic mortality increased following APAP exposure as determined by an increased number of resorption points as compared to the control group (Fig. 4b-c, n=19-20). These in vivo results show that exposure to APAP impacts the ability of embryos to implant and/or survive after implantation.

APAP reaches the human reproductive organs and disrupt human development at therapeutic doses

To understand to what degree APAP enters the reproductive organs in women, follicular fluid (n=26), endometrial tissue (n=7), and uterine fluid (n=7) were collected as part of routine procedures in the clinic from patients receiving a standard therapeutic dose of 1g of APAP. 1h following administration, APAP had reached the follicular fluid at an average concentration of 38.1 μM, the endometrial tissue at an average of 80.3 μM, and the uterine fluid at an average of 124.5 μM. Notably, one woman had 291.3 μM in the uterine fluid (Fig. 5a).

To investigate the effect of APAP on human PID at concentrations present in the intrauterine environment after standard therapeutic doses, we exposed human cleavage stage embryos to 100 and 200 µM APAP. For the exposure to 100 µM APAP, human development day 2 (D2) and D3 embryos (n=6) were cultured for 48 or 72 h to reach D5 with corresponding controls (n=6). For the exposure to 200 μM APAP, D2 embryos (n=5) were cultured for 72 h to reach D5 with corresponding controls (n=5). Results indicated that embryonic development in the APAP groups were compromised as compared to most of the controls (Supplemental video 1 and 2; Fig. 5b, c - top panel; Supplemental Fig. 3b).

In embryos exposed to 100 µM APAP, the overall structural organisation of the embryos appeared normal, but development was delayed as enumerated by reduced cell numbers as compared with controls (Fig. 5c - middle and lower panels). The concentration of 200 µM resulted in amorphous cells with fragmented nuclei indicating cell death (Fig. 5b - middle and lower panels). Quantifying the DAPI nucleus staining from each experiment, we observed a reduction in cell number of both 100 and 200 μM exposed embryos as compared to control (Fig. 5d-e). These data show that at concentrations present in the intrauterine environment after a standard therapeutic dose, APAP disrupt early human PID either by delaying development or direct embryonic fatality.

APAP exposure inhibits DNA synthesis in human blastocyst stage embryos

To investigate the effect of APAP on DNA synthesis in situ in late human PID, blastocysts stage embryos D5-6 were exposed to APAP for 6 h (100 and 200 μM) with thymidine analogue EdU (5-ethynyl-2´-deoxyuridine) added for the last 3 h (Fig. 6a, e) to quantify DNA synthesis. The percentage of APAP-exposed embryos with a definable ICM was reduced as compared to controls (n=14-18) at a dose of 100 µM (n=18) with a similar tendency observed with 200 µM (n=15) (Fig. 6b, f).

Finally, we investigated DNA synthesis of OCT3/4 positive cells in the embryos with a definable ICM. We found no difference in number of OCT3/4 positive cells with incorporated EdU between embryos exposed to 100 μM APAP (n=11) as compared with controls (n=18) (6c). However, we found that the level of EdU incorporation was reduced in the OCT3/4 positive ICM cells as compared with controls (Fig. 6d: control; n=108 (108 cells from 18 blastocysts), APAP; n=60 (60 cells from 11 blastocysts)). In the embryos subjected to 200 µM APAP (n=9), the percentage of OCT3/4 positive cells with EdU incorporation was decreased (n=12) (Fig. 6g). As with the 100 µM APAP exposure blastocyst, we found the level of EdU incorporation was reduced in the OCT3/4 positive ICM cells as compared to controls (Fig. 6h: control; n=71 (71 cells from 11 blastocysts), APAP; n=37 (37 cells from 9 blastocysts)). Taken together, these results suggest that APAP exposure for 6 h disrupt human PID at the blastocyst stage at physiological relevant concentrations through inhibition of DNA synthesis and proper expansion of ICM cells (for raw data see Supplemental Table 1a and 1b).

The first days during PID are perhaps the most critical during human development^6,7. It is estimated that 10-40% of early embryos are lost before or at the time of implantation^3–7,38. We here show that APAP disrupts embryonic development during this sensitive period of life by inhibiting the cell cycle. Although we cannot rule out effects involving pharmaceutic targets such as PTGS1, PTGS2 and CB1, the data from S. pombe point towards inhibition of RNR as a plausible cause. Consistently, we observed DNA synthesis inhibition in in vitro models, such as human embryonic stem cell lines, and in vivo in human blastocyst stage embryos.

Using mouse embryos as a model, we found a time and concentration dependent effect on growth and the rate of blastocyst formation of embryos exposed to >25 µM APAP. Following transfer of embryos to recipient pseudo-pregnant dams, the number of implantations and full-term fetuses was decreased in uterine horns with transferred APAP exposed embryos as compared to uterine horns with transferred control embryos. These data indicate that disruption of PID by APAP exposure for 24 or 48 h reduced the chances of development to term.

Our studies indicate that human and mouse cleavage stage embryos were more sensitive to APAP as compared to the blastocyst stage, resulting in embryonic fatality at a concentration of 200 µM. Nonetheless, in the mouse we also observed an effect on the ICM of blastocysts stage embryos at 200 µM. A similar effect was observed in human blastocyst stage embryos with 100 µM after 6 h with a comparable tendency at 200 µM. This suggests not only that the pluripotent ICM resembles the blastomeres of cleavage stage embryos in sensitivity to APAP, but also that only a few hours of exposure are enough to cause negative effects on PID. Importantly, these concentrations are similar or lower than what we found in the reproductive organs including the uterus of women after a single therapeutic dose of APAP where the maximal concentration identified was 291.3 μM.

In our mouse pregnancy model, we found effects on both the number of offspring and implantations after gavage with 200 mg/kg/day. As the maximal human dose is 50 mg/kg/day, this is below the safety margin of 12.33x used when translating preclinical doses from mouse to human³⁹. Implantation is dependent on a timed interaction between the developing embryo and the hormonally primed endometrium. For successful implantation in humans, embryos must progress to the blastocyst stage and undergo timely hatching⁵. Delayed development results in asynchrony between the developing embryo and the endometrium, increasing the risk of implantation failure or miscarriage⁵. Moreover, ICM quality, defined by cell number and morphology, has been shown to be the strongest predictor of live birth following embryo transfer⁴⁰.

Based on the present data, we suggest that APAP (depending on timing, duration and concentration) disrupts early development in three different scenarios: (i) direct embryonic fatality at higher doses ≥200 µM, (ii) failed implantation due to asynchrony of the embryo and endometrium by delayed PID, and (iii) miscarriage after implantation due to disruption of PID in situ e.g. reduced cell number in ICM.

As early embryonic fatality or failure of implantation cannot be easily identified in human, they are often interpreted as subfertility by health professionals⁶. However, the impact of APAP on overall fertility might be substantial as APAP is one of the most frequently used medications globally²² and has long been considered as a safe option for treatment of pain and fever during pregnancy by regulatory bodies such as the FDA and EMA when used as directed^23,24. Moreover, several studies from Europe have shown that all citizens are exposed to APAP from the environment^12–14, indicating that APAP is not only a pharmaceutical but also a significant environmental pollutant.

A limitation of the present data is the number of donated human cleavage stage embryos. Our translational approach with reproducibility across multiple model systems circumvent this limitation to some extent, e.g., by the recapitulation of cell cycle effects from yeast to mouse preclinical models and human embryos. Further studies are now warranted to substantiate these data, including investigating APAP use among fertile healthy women and the possible link to pregnancy loss in prospective cohorts. As cell division is at the heart of all development, more studies are also needed to understand how environmentally caused cell cycle inhibition can affect other processes during development.

APAP concentrations in female reproductive tract

All materials were sampled at the Copenhagen University Hospital, Denmark, and TFP Stork Fertility clinic, Denmark, in accordance with relevant guidelines and regulations and after consent from the regional scientific ethical committee of the Capital Region of Denmark (protocol nr.: 17003845). All material was anonymised and collected as part of standard clinical procedures.

Follicular fluid: Follicular fluid was collected from a total of 26 women during transvaginal ultrasound guided aspiration of the pre-ovulatory follicles. The procedure is part of standard fertility treatment with the aim of collecting oocytes, termed oocyte pickup (OPU). As a by-product of OPU, follicular fluid is recovered but is routinely discarded. Each patient scheduled for OPU receive, as part of pain-management 1 gram of APAP 1 hr before OPU. Collected follicular fluids were centrifuged and stored at -20°C until analysis.

Endometrial tissue and uterine fluid: Endometrial tissue and uterine fluid were collected from 7 women as part of an endometrial scratching (ES) procedure. ES is performed as part of fertility treatment to patients with recurrent implantation failure and has been suggested to improve chances of implantation^41,42. Endometrial tissue and uterine fluid are recovered during ES but is routinely discarded. Each patient scheduled for ES received as part of pain-management 1 gram of APAP 1 hr before the procedure. Following intrauterine placement of an inner and outer biopsy catheter a small amount of suction was applied. The inner catheter was removed and brought to the laboratory for collection of uterine fluid. A new inner catheter was then positioned, and endometrial biopsy/scratching was performed by moving the catheter while applying strong suction with a syringe. In the laboratory the initial inner catheter was rinsed with 0.4 ml of sterile saline to recover uterine fluid. Fluid from the first inner catheter and endometrial tissue from the second catheter was stored at -20°C until analysis.

Liquid chromatography–mass spectrometry (LC-MS/MS) measurements for APAP: For measurements of APAP in follicular fluid, 200 uL aliquots of follicular fluid, calibration standards (10 solutions of native APAP standard diluted in water in the concentration range from 0.5-2000 ng/mL) and control materials (native APAP spiked in urine pool in three different concentrations) were added 20 µL of internal standard solution followed by 276 µL NH₄Ac-buffer. Immediately before enzymatic de-conjugation, all sample extracts calibration and control materials were added 10 µL freshly prepared enzyme mixture (ß-glucuronidase from Escherichia coli K12), sulfatase from Aerobacter aerogenes, and NH₄Ac-buffer; 1:1:3), mixed and incubated at 37ºC for 3 h, stored overnight at -20ºC and then centrifugated at 4ºC for 10 min. Supernatants were transferred to HPLC vials and were then ready for analysis.

For uterine fluid and endometrial tissues, APAP were extracted from approximately 100 mg of uterine fluid (98.9-120 mg) and endometrial tissue (35.2-141 mg) samples following a validated method for extraction of chemicals⁴³. Briefly, samples were added 20 µL of internal standard solution (containing 200 ng/mL of APAP-d4 and 100 ng/mL ¹³C₄-methylumbelliferone including 4-methylumbelliferyl b-D-glucuronide and 4-methylumbelliferyl-b-D-sulfate dissolved in 50% methanol), then centrifuged and stored at RT for 30 min. Samples were submerged by addition of 1 mL acetone and mechanically homogenized with a mixer. The mixer was then washed with 2 mL methanol per extract, which was collected and added to the homogenized extract. Then the extracts were sonicated in an ultrasound bath for 10 min. Subsequently, total extract volumes were reduced to < 2 mL each by evaporation under a gentle nitrogen stream at RT. Thereafter tissue residuals were removed by transferring the remaining extracts to a 2 mL Eppendorf tubes followed by storing at -20ºC for 15 min and then centrifugation at 4ºC for 10 min. Supernatants were then transferred to new glass tubes, evaporated to dryness under a stream of nitrogen and re-suspended in 496 µL 0.5 M ammonium acetate (NH₄Ac) buffer (pH5.5).

The total (free and conjugated) content of APAP in the sample extracts were measured by isotope diluted online-TurboFlow-liquid chromatography mass spectrometry (LC-MS/MS) using a Thermo Scientific Aria TLX-1 LC system coupled to a TSQ Ultra triple quadrupole mass spectrometer equipped with a heated electrospray ionization source (HESI) running in positive mode. The instrument was used in combination with Aria operating software 1.6.3 and Xcalibur 2.1.0.1139 system software (ThermoFinnigan, Bellefonte, PA, USA). The TurboFlow-LC systems were equipped with a loading column; TurboFlow Cyclone P column, 0.5 x 50mm (Thermo Scientific) followed by an analytical Gemini-C18 column, 3 µm particle size, 3 x 50 mm (Phenomenex). Prepared batches were kept on the auto-samplers at 10°C. The injection volume was 100 µL. Flow rate and loading and eluting gradients were specified for this specific method and the mobile phases used were loading solvents; A: 10 nM NH4AC, B: 0.1% formic acid in methanol, C: acetone/isopropanol/acetonitrile 10:45:45 and eluting solvents; A: 3 nM ammonium hydrogen carbonate, B: acetonitrile. The method was validated, and limit of detection determined (LOD = 0.48 ng/mL) for urinary APAP as previously described⁴⁴ according to the ICH guidelines⁴⁵. The sample extracts were analysed in one batch, also including standards for calibration curves, three blanks and three times three spiked urine pool controls followed by one batch more with samples diluted to fit the method calibration range. The relative standard deviation (RSD) in the three control levels ranged from 2.7-7.6%. For the LC-MS/MS analyses, native APAP (N-acetyl-4-aminophenol, CAS No. 103-90-2 procured from Sigma-Aldrich), labelled APAP-d4 (Paracetamol-D₄, CAS No. 64315-36-2 procured from LoGiCal^®) and all other reagents and solvents were of analytical, HPLC or MS grade, and all chemicals and laboratory equipment were tested for contamination before utilisation.

Human embryogenesis experiments

All experiments were performed in accordance with relevant guidelines and regulations and conducted at the Copenhagen University Hospital – Hvidovre and Copenhagen University Hospital - Rigshospitalet, Denmark. Surplus human embryos from fertility treatment were donated at Copenhagen University Hospital - Rigshospitalet, Copenhagen University Hospital - Hvidovre, TFP Stork Fertility Clinic, and Copenhagen Fertility Center between the years 2020-2023. The study protocol was approved by the Research Ethics Committee of the Capital Region of Denmark (H-19050437) with signed informed patient consent given prior to donation. A total of 22 cleavage stage embryos and 68 blastocyst stage embryos were used. All experiments were performed with APAP dissolved directly in culture media without any changes in osmolarity (data not shown). APAP was purchased from Sigma cat. no A5000 (Sigma/Merck, Darmstadt, Germany) for these experiments and all subsequent experiments in the study.

Cleavage stage embryos: Cleavage stage embryos were frozen at D2 or D3 after fertilisation and thawed using a slow freeze protocol (Freezekit and ThawKit Cleave Vitrolife, Göteborg, Sweden) as described by the manufacturer (https://www.vitrolife.com/globalassets/support-documents/short-protocols/sp_slow_freeze_cryopreservation_Cleavage.pdf). Following thawing, embryos were placed in equilibrated SAGE 1-Step media (CooperSurgical Fertility Solutions, Ballerup, Denmark) drops covered with mineral oil (Origio, CooperSurgical Fertility Solutions, Ballerup, Denmark) at 37 ºC under 6 % CO₂ and 5 % O₂. Within 2 h after thawing embryos were matched in pairs and allocated to exposure groups (100 or 200 µM APAP) or control group according to a prioritized order of the following parameters: (i) female origin (sibling embryos), (ii) female age at time of embryo cryopreservation, (iii) time of cryopreservation (D2 or D3), (iv) and number of blastomeres and morphology after thawing. This was done to minimize differences between control and APAP exposed blastocysts. Experiments were performed using a timelapse incubator (EmbryoScope ESD Vitrolife, Göteborg, Sweden). As the specific cryopreservation time of the individual D2 embryos were unknown, the start point depicted in time-lapse videos (Supplemental video 1 and 2) were estimates of embryo age after fertilisation.

Blastocyst stage embryos: Embryos were vitrified at D5 or D6 and subsequent thawed (Vit Kit Freeze NX and Vit Kit®-Warm, Irvine Scientific, Santa Ana, CA, USA) as described by the manufacture (https://www.irvinesci.com/media/IrvineScientific/Resources/0/0/002773_warming_ooctyes_protocol.pdf). Embryos were subsequently transferred to a pre-equilibrated dish with 25 µl drops of SAGE 1-Step culture media (CooperSurgical Fertility Solutions, Ballerup, Denmark) and cultured at 37 ºC under 6 % CO₂ and 5 % O₂. Within 3 h after warming embryos were matched in pairs and allocated to exposure groups (100 or 200 µM APAP) or control group according to a prioritized order of the following parameters: female origin (sibling blastocysts), female age at time of vitrification, day of vitrification (D5 or D6) and morphology post warming. A maximum of 12 embryos were warmed per experiment by two experienced embryologists to reduce difference in culture time between warming and experiment initiation. Morphology was evaluated by light microscopy (LM) following warming, at start of the experiment, following 3 h of APAP exposure, and at the end of the 6 h culture period using the Gardner grading system⁴⁶ (Supplemental. Table 1a and 1b).

Whole-mount immunofluorescence staining of human blastocysts stage embryos: Click-iT^® Plus EdU Alexa Fluor^® 555 Cell Proliferation imaging kit (Thermo Fisher Scientific, Waltham, MA, USA) was used to visualize and quantify newly synthesized DNA. APAP was diluted in equilibrated SAGE 1-Step media and dishes made with 25 µl culture media drops with or without APAP (100 or 200 µM) covered in mineral oil (Origio, CooperSurgical Fertility Solutions, Ballerup, Denmark) and maintained in the incubator for minimum 2 h before experiment start. Embryos were exposed to 100 or 200 µM APAP and control in SAGE 1-Step media for 6 h with the addition of 10µM EdU for the final 3 h. After treatment, the embryos were subjected to a modified procedure of the protocol from Wong (2021)⁴⁷ combined with the manufacture protocol of the Click-iT^® Plus EdU Alexa Fluor^® 555 Cell Proliferation imaging kit. In brief, Embryos were fixed individually in Millicell® (Sigma/Merck, Darmstadt, Germany) cell culture inserts in 4% paraformaldehyde (PFA) (VWR chemicals, Radnor, PA, USA) for 15 minutes at room temperature (RT). Next, the embryos were washed twice in 3% BSA in PBS before placed in 0.5% PBSTr for 20 minutes at RT. Again, the embryos were washed twice in 3% BSA in PBS and placed in Click-iT® Plus reaction cocktail for 30 minutes. The embryos were protected from light for the remainder of the protocol and were washed once in 3% BSA in PBS before transferred to neutralization buffer and incubated for 15 minutes at RT. Embryos were then subjected to washes, primary- and secondary antibodies, and mounted as described under “Whole-mount immunofluorescence staining of human and mouse embryos”. Segregation between the ICM and TE fates was determined by the expression of OCT3/4 and CDX2^37,48. A definable ICM was identified by a clear cluster of cells exclusively expressing OCT3/4. An indefinable ICM was defined by either no OCT3/4 positive cells, OCT3/4 positive cells dispersed throughout the blastocyst stage embryos with no cluster or a cluster of cells expressing both CDX2 and OCT3/4.

Whole-mount immunofluorescence staining of human and mouse embryos: The whole-mount immunofluorescence staining of embryos was performed following a procedure modified from Wong 2021⁴⁷. In brief, preimplantation embryos were fixed in 4% paraformaldehyde (PFA) (VWR chemicals, Radnor, PA, USA) for 15 minutes at RT with human embryos fixed individually using Millicell® (Sigma/Merck, Darmstadt, Germany) cell culture inserts. Next, the embryos were placed in 0.5% PBSTr (0.5% Triton X-100 (Sigma/Merck, Darmstadt, Germany) in PBS) for 20 minutes at RT and then transferred to neutralization buffer (1M Glycine in 0.1% PBSTr) and incubated for 15 minutes at RT. The embryos were then washed three times in 0.01% PBSTw (0.01% Tween-20 (Sigma/Merck, Darmstadt, Germany) in PBS) for 10 minutes at RT. After the last wash, the embryos were transferred to a well containing blocking buffer (3% donkey serum and 1% BSA (Sigma/Merck, Darmstadt, Germany) in PBSTw) and incubated in a humidified box containing wet tissue paper at 4°C overnight. The embryos were then transferred to primary antibodies diluted 1:200 in blocking buffer and incubated for 2-3 days at 4°C. The embryos were then washed 3 times in 0.01% PBSTw for 10 minutes at RT. The embryos were then transferred to a well with secondary antibodies diluted 1:500 in blocking buffer (DAPI diluted 1:5,000 or phalloidin diluted 1:100) and incubated, in the dark, at RT for 3-4 h. Afterwards, the embryos were washed 3 times in 0.01% PBSTw for 10 minutes at RT before being transferred to a coated microscope slide (Dako Agilent, Glostrup, Denmark). The slides were left to dry for a few minutes at RT, in the dark, before addition of 10 μl of mounting medium (90% glycerol and 2% n-propyl gallate in PBS) and mounting with cover glasses (VWR, Radnor, PA, USA)⁴⁹. The cover glass edges were sealed with transparent nail polish.

Immunofluorescence microscopy and imaging: The immunofluorescence and differential interference contrast (DIC) images were obtained using an Olympus BX63 upright microscope with an Olympus DP72 color, 12.8-megapixel, 4140 Å~3096 resolution camera (Olympus, Tokyo, Japan). Olympus CellSense Dimension software version 1.7 (Olympus, Tokyo, Japan) was used for deconvolution and 3D reconstruction of Z-stacks as previously described⁵⁰. Images were later processed in ImageJ (Bethesda, MD, USA) and Adobe Photoshop CS6 (Adobe, San Jose, CA, USA). For quantification of the EdU intensity, an outline was drawn around each cell double positive for OCT3/4 and EdU. Only cells or parts of cells that did not overlap with other cells were included. Using the measurement and region of interest (ROI)-function in the CellSens dimension software (Olympus, Tokyo, Japan), the mean red fluorescence intensity was measured along with a background reading. The corrected mean fluorescence for the individual cells were subsequently calculated by subtracting the corresponding background value. The following primary antibodies were used at a 1:200 dilution: rabbit anti-CDX2, #D11D10 (Cell Signaling Technology, Danvers, MA, USA), goat anti-OCT3/4, #sc-8629 (Santa Cruz Biotechnology, Dallas, TX, USA), mouse-GATA4, #sc-25310 (Santa Cruz Biotechnology, Dallas, Tx, USA), and mouse anti-α-Tubulin, #T5168 (Merck/Sigma-Aldrich, Darmstadt, Germany). Secondary antibodies (all from Thermo Fisher Scientific, Waltham, MA, USA) were used at a 1:500 dilution: Donkey-anti-mouse IgG Alexa Fluor® 568, Donkey-anti-rabbit IgG Alexa Fluor® 568, Donkey-anti-goat IgG Alexa Fluor® 488, and Donkey-anti-rabbit IgG Alexa Fluor® 647. F-actin and nuclei were stained with Phalloidin (Alexa Fluor® 488 or 568, Thermo Fisher Scientific, Waltham, MA, USA) and DAPI (Thermo Fisher Scientific, Waltham, MA, USA), respectively.

Mouse models

Mouse pregnancy model: Experiments were performed under license number 2019-15-0201-00175 from the Danish Animal Experiments Inspectorate and under EU directive 2010/63/EU on the protection of animals used for scientific purposes. C57Bl6/J BomTac mice (Taconic, Lille Skensved, Denmark) arrived at the animal facility at 7-8 weeks of age. Experiments were run in two independent cohorts of 24 females and 12 males. Upon arrival, females were housed in twelve boxes of two and males six-by-six. Otherwise, housing and environment was as described previously⁵¹. Mice were acclimatized 2 weeks prior to starting the experiments. On the day of experiment initiation, one male was introduced to two dams in one home cage and the following day (day 1) APAP administration was initiated. Female cages were allocated by draw to either 200 mg/kg/day APAP administrated as a single daily oral gavage in tap water or tap water (vehicle control) for a total of 10 days (days 1-10). Females sharing cages received the same treatment and dosing based on body weight on day 0. Males were separated from females on day 5 to avoid further breeding. After end administration at day 10, dams were transferred to clean cages. Dams were euthanized on day 18 and number of fetuses and resorptions counted blindly by two experienced veterinarians. 3 controls and 4 APAP dams were excluded due to lack of pregnancy.

Mouse embryo culture and transfer model: Experiments were performed under license number 2021-15-0201-00851 from the Danish Animal Experiments Inspectorate and under EU directive 2010/63/EU on the protection of animals used for scientific purposes. For the mouse embryo collection, culture, and embryo transfer, inbred C57BL/6JRj females were used as embryo donors and outbred CD1 (RjOrl:SWISS) were used as embryo recipients. Mice were kept in individually ventilated cages at a temperature of 22 °C (±2 °C), with a humidity of 55% (±10%), under 12/12-hr light/dark cycles. Embryos were harvested from prepubescent (4–5-week-old) C57BL/6JRj females. The donor females were subjected to a hormone treatment before mating that consisted of an intraperitoneal injection of 5 IU/female of PMSG (Pregnant Mare Serum Gonadotropin, Prospec, Rehovot, Israel,), followed 47 h later by a second intraperitoneal injection of 5 IU/female of hCG (Human Chorionic Gonadotropin, Chorulon Vet, ref. 422741). After the second injection, each female was set in cross with a C57BL/6JRj stud male. The following morning, mating was monitored by the observation of copulation plugs in the vagina of the females. The day of plug detection was considered embryonic day 0.5 (E0.5). 2-cell-stage embryos (E1.5) were harvested 1.5 and morulae 2.5 dpc. On the day of dissection, pregnant females were sacrificed by cervical dislocation and immediately after the oviducts were dissected and placed on a petri dish containing M2 medium (Sigma/Merck, Darmstadt, Germany, ref. M7167). Embryos were flushed out of the oviduct using a blunt 30G needle attached to a 1 ml syringe filled with M2 medium. Embryos were collected from the eluted medium using a 115 μm diameter glass capillary (Retransferpipette, Biomedical Instruments, Zöllnitz, Germany) attached to a mouth-pipetting system (Mouthpipette, Biomedical Instruments, Zöllnitz, Germany) and washed through three 50 μl drops of M2 medium. Finally, embryos were moved to a fresh petri dish and placed in a 50 µl drop of KSOM medium (Embyomax KSOM + AA, Merck Millipore, Burlington, MA, US, ref. MR-106-D) covered by sterile mineral oil (NidOil, NordicCell, Copenhagen, Denmark, ref. 90142). Embryos were cultured for either 24 or 48 h at 37C, 5% C0₂, in KSOM only (control groups) or in different concentrations of APAP diluted in KSOM. All experiments were performed with APAP dissolved directly in culture media without any changes in osmolarity (data not shown).

For embryo transfer to pseudo pregnant recipient dams, 10–12-week-old CD1 females in anestrus were stimulated to enter the cycle by setting them in cross with vasectomized CD1 males. Formation of vaginal plug was monitored every morning for three days. Dams that showed a plug on the third day of breeding were selected as pseudo pregnant females and used to perform the embryo transfer of the cultured embryos. The dam was anesthetized by an intraperitoneal injection of 20 mg/ml Avertin from a stock solution of 1g Tribromoethanol (VWR chemicals, Radnor, PA, USA, ref. ACRO421430500) diluted in 630 µl 2-methyl-2-butanol (Sigma/Merck, Darmstadt, Germany, ref. 240486). 120 µl of this stock was diluted up to 10 ml in saline water to obtain the ready-to use solution at a dose of 25 µl/gram. 10-12 embryos were transferred to each oviduct of the pseudo pregnant dam. The embryo transfer was performed according to the standard procedure described in⁵². Control embryos were always transferred to the right oviduct, and APAP exposed embryos to the left oviduct. 18 days after the transfer (embryonic day E18.5), the pregnant dams were euthanized and dissected to count the number of fetuses developed in each uterine horn.

Human cell culture

Cell culture: HEK293 cells were cultured in high glucose Dulbecco's modified Eagle's medium (DMEM, Thermo Fisher Scientific, Waltham, MA, USA) supplemented with 10% fetal bovine serum (FBS, Thermo Fisher Scientific, Waltham, MA, USA) and 1% penicillin–streptomycin. The cells were kept at 37°C in a humidified 5% CO₂ incubator and cultures beyond passage 25 were discarded. The human embryonic stem cell lines H1 (WA01, WiCell; RRID: CVCL_9771) and PDX1EGFP/+ HUES4, acquired from our facility⁵³, were cultured in DEF-CS culture media (Takara Biosciences, San Jose, CA, USA) according to the manufacturer's guidelines. The culture medium was changed daily, and cells were passaged every 2-3 days using TrypLE Express Enzyme (Thermo Fisher, Waltham, MA, USA). All cell cultures were maintained in a humidified incubator at 37°C with 5% CO₂. Culturing the human embryonic stem cell lines H1 and HUES4 was done in accordance with relevant guidelines and regulations and after consent from the regional scientific ethical committee of the Capital Region of Denmark (protocol nr.: H-21043866; appendix 94634).

Cell- and viability count: HEK293 cells were plated in 60 mm petri dishes (≈ 20% confluency) and HuES4 and H1 cells were seeded in 12-well cell culture dishes with a density of 15,000 cells per well. After 24 h of settling, cells were treated with either DMSO (control) (Sigma/Merck, Darmstadt, Germany) or APAP (HEK293: 500 µM, HuES4 and H1: 200 µM), for 24, 48, or 72 h. To detach the cells, Trypsin or TrypLE Express Enzyme (Thermo Fisher, Waltham, MA, USA) were used. The cells were then centrifuged at 1000 rpm for 5 minutes, resuspended in 1 mL of culture medium, and counted. HuES4 and H1 cell counting was performed on quadruplicate technical replicate wells. The counting process was performed with Via1-Cassettes™ (ChemoMetec, Lillerød, Denmark) and a NucleoCounter® NC-200™ (ChemoMetec, Lillerød, Denmark) automated cell counter following the manufacturer's protocol and resulted in measurement of total number of cells and the percentage of viable cells. All three cell lines were analysed at a ≈ 80 % confluency to avoid restricted growth and to ensure that the analyses was performed on cells within their exponential growth phase.

Flow cytometric analysis of HEK293, HuES4, and H1 cell lines: HEK293, HuES4, and H1 cell lines were plated and left to settle for 24 h prior to treatment with control (DMSO) or APAP for 48 h. The cells were detached, fixed by slowly adding ice cold 70% ethanol while gently vortexing the tube, and left overnight in the fridge at 4°C. After centrifugation and removal of the supernatant, the pellet was resuspended in PBS containing 0.25% triton X-100 and incubated on ice for 15 min to permeabilize the cells. The cell pellet was then incubated for 30 min in the dark at RT with PBS containing 10 ug/mL RNase A (diluted from a stock 10 mg/mL RNase A (Sigma/Merck (R-5000), Darmstadt, Germany) in 10 mM Tris pH 7.5, 10 mM MgCl₂) and 20 ug/ml propidium iodide (diluted from a stock 3.6 mg/ml propidium iodide (Sigma/Merck, Darmstadt, Germany) in DMSO). After DNA staining, the cells were loaded and analysed on a FACSverse multicolour flow cytometer using the FACSuite software (BD Bioscience, San Jose, CA, USA). Cells were gated according to forward-scatter/side-scatter (FSC/SSC) principles, followed by an FSC-A/FSC-H to ensure the analysis of single cells. Approximately 6.000-10.000 cells were analysed per condition. For an unbiased analysis of the cell cycle profile the Flowing Software 2.5 (Turku Bioscience Centre, Turku, Finland) automated cell cycle tool was used to define the distribution of cells within the different cell cycle stages, G1/G0, S, and G2/M. All three cell lines were harvested at a 70-80 % confluency to avoid restricted growth and to ensure that the analyses were perform on cells within their exponential growth phase.

DNA replication analysis: Click-iT^® Plus EdU Pacific Blue^® Flow Cytometry Assay Kits (Thermo Fisher Scientific,Waltham, MA, USA) were used to measure de novo DNA synthesis according to the manufacturers protocol. Briefly, HEK293, HuES4, and H1 cell lines were plated and left to settle for 24 h prior to treatment with control (DMSO) or APAP for 3 h with the addition of 10µM EdU for the final 2 h. Cells were loaded and analysed on a FACSverse multicolour flow cytometer using the FACSuite software (BD Bioscience, San Jose, CA, USA). Cells were gated according to forward-scatter/side-scatter (FSC/SSC) principles, followed by an FSC-A/FSC-H to ensure the analysis of single cells. Approximately 6.000-10.000 cells were analysed per condition. For the detection of EdU Pacific Blue^® a 405 nm excitation filter with a violet emission filter (448/45) were used.

Fission yeast culture

For growth assessment on solid agar, fission yeast (S. pombe) strains of indicated genotypes were grown over night in yeast extract liquid medium. Cells were then counted, and diluted to 5000, 500, 50 and 5 cells/µl. 5 µl of each dilution were spotted onto rich medium agar plates containing 1% DMSO or 1% DMSO/APAP and incubated for 3 days at 32°C before photography. For growth assessment in liquid yeast extract, fission yeast strains of indicated genotypes were grown at 32°C in the presence of 1% DMSO or 40 mM APAP/1% DMSO and were monitored by triplicate cell counting using a NucleoCounter® NC-3000 ™ (ChemoMetec, Lillerød, Denmark) as described by the manufacturer. Doubling time was calculated assuming exponential growth. Samples for DNA content were fixed in 70% ethanol, washed in 20 mM EDTA, and treated with RNAse A overnight before staining of the DNA with CytoxGreen and analysis of cellular DNA content using the NucleoCounter® NC-3000 ™ (ChemoMetec, Lillerød, Denmark) as described by the manufacturer.

Data analysis and statistics

Data analyses were performed with GraphPad Prism 9 (San Diego, CA, USA), except for the Fisher-Freeman-Halton and Phi and Cramer's tests that were performed in SPSS 29.0.1.0 (Chicago, IL, USA). For the cell cultures the ‘n’ signifies biological replicate experiments obtained from different passages. For mouse and human embryo culture experiments individual embryo was considered as one n. For animal studies each animal was considered as one n. The specific statistical test used to analyse the given data is stated in the figure legends. Data is represented as the mean ± SD or SEM and P< 0.05 was considered statistically significant.

Acknowledgements

This work was supported by the Lundbeck Foundation (R324-2019-1881). Authors P.S. and H.K.M. are part of the Novo Nordisk Foundation Center for Stem Cell Medicine (reNEW; NNF21CC0073729).

Competing interests

Authors declare no competing interests.

Author contributions

D.M.K., B.S.N., K.K., and M.R.P. conceived and designed the study. D.M.K., B.S.N., M.R.P, J.M., C.H., H.K.M., H.F., C.R., E.M.J., P.S., S.L.T.C., N.R.J., A.H., P.A.P., A.J., A.P., S.Z., J.E., A.Z., and S.T.C. performed experiments. D.M.K., B.S.N., M.R.P., K.B.P., S.L., J.E., F.L., and A.Z. coordinated the surplus human embryo donations. D.M.K., B.S.N., M.R.P., and S.T.C. analysed and interpreted the data. Drafting the article was performed by D.M.K., B.S.N., and M.R.P. All authors have read and approved the final version of this manuscript.

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Human cleavage stage embryos exposed to 100 µM paracetamol
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Human cleavage stage embryos exposed to 200 µM paracetamol
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Paracetamol (N-acetyl-para-aminophenol) disrupts early human embryogenesis

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