Identification of a novel natural compound that acts on the membrane progestin receptor α (paqr7) from the marine algae Padina.

All the methods in this study were performed in accordance with the ARRIVE guidelines 34 . The use of zebrafish, mice and the experimental protocol were approved (approval no. 2024F-9, 2024A-14) by the Institutional Ethics Committee of Shizuoka University, Japan. All procedures were performed in accordance with the relevant institutional guidelines and regulations. The following chemicals and materials were purchased: an ODS-SM column (50 μm, 1.8 × 11.4 cm, 7 g; Yamazen Corporation, Japan); hyaluronidase from bovine testes (Tokyo Chemical Industry, Japan); HCG (Mochida Pharmaceutical Corporation, Tokyo, Japan); and PMSG (Asuka Animal Health Co., Ltd., Tokyo, Japan). The marine algae Padina arborescens was collected from the Mochimune Marine Field of the Faculty of Agriculture, Regional Field Science Education and Research Center, Shizuoka University (34°92′07.51’'N, 138°36′82.14" E). Species identification was performed by DNA sequencing as previously described 22 . The sample was washed with tap water and dried in the sun. The dried sample was then powdered using a ball mill (Retsch PM-100) under cryogenic conditions with liquid N 2 . Then, 150 g of powdered Padina was soaked in 1500 mL of 100% methanol for 7 days with occasional stirring. The soluble component was separated from the insoluble portion using Whatman filter paper. The filtrate was solidified using a rotary evaporator (As One; ARE-V1200) operated at 40 °C and stored at 4 °C. The actual yield from this step was 14.7%. Primary purification of the crude methanolic extracts was performed by separating the polar and nonpolar compounds via liquid‒liquid extraction (80% aqueous methanol:n-hexane = 1:1) using a separating funnel and concentrating with a rotary evaporator. The percentage yield of polar compounds from this step was 3.692%. In the next step, the solid polar fraction of the extracts was redissolved in 100% EtOH, diluted tenfold with ultrapure water (DDW), and applied to an ODS-SM column (50 μm, 1.8 × 11.4 cm, 7 g, Yamazen Corporation, Japan) connected to a peristaltic pump. All the compounds in the polar fraction were loaded onto fifty ODS-SM columns and fractionated by HPLC using a gradient elution program of acetonitrile (0–100%) and water with 0.05% trifluoroacetic acid (TFA). Eight different colored fractions were separated, including the whole of the ODS-SM column. The separated fractions were concentrated by lyophilization using a freeze dryer (FDU-810, EYELA, Tokyo, Japan) and redissolved in 100% EtOH for further purification. Then, three consecutive HPLC purification steps were performed using three different stationary phase columns to obtain a single purified compound peak (Supplementary Fig. S1 ). The target fractions from the ODS-SM column were further fractionated by the HPLC separation technique using silica-based C18 columns (10 × 300 mm) as the stationary phase and a linear gradient of acetonitrile (0–100%) under acidic conditions (0.05% TFA) as the mobile phase. The target mPRα-interacting fraction from the C18 column eluate was further purified via the TSKgel Phenyl-5PW RP glass column (8 mm × 7.5 cm). Finally, a single peak compound with mPRα-interacting activity was purified using a Wakopak Navi C30-5 column (2.0 × 150 mm) with a gradient elution of acetonitrile (0–100%) maintained under slightly acidic conditions with 0.05% TFA. The UV detector was operated at 215 nm for all steps, and the column oven was maintained at 40 °C. The fractions were collected at room temperature and concentrated by lyophilization. The dried samples were redissolved in 100% EtOH and stored at 4 °C for further analysis. The percentage yield of the purified compound after final purification was 0.0033%. The final purified single peak obtained from the HPLC purifications was analyzed by NMR. The purified fraction was lyophilized and then dissolved in 500 µL of CDCl 3 . The solvent impurity was used as a reference ( 1 H: 7.26, 13 C: 77.36). The NMR spectra, including 1 H, 13 C, DQF-COSY, TOCSY, HMQC and HMBC, were measured using a JNM-ECZ500R spectrometer (JEOL, Tokyo, Japan) according to the manufacturer’s instructions. The LC–MS conditions were as follows: LC, Accela pump (Thermo Fisher Scientific, Waltham, MA, USA); column: CAPCELL PAK C18 MGIII (150 × 4.6 mm, 3 μm; Osaka Soda, Osaka, Japan); solvent: 0.1% formic acid (A) and acetonitrile (B); elution: 0–2 min, isocratic at 5% B; 2–23 min, linear gradient from 5 to 95% B; 23–25 min, isocratic at 95% B; flow rate: 0.35 mL/min; MS: Orbitrap Exactive (Thermo Fisher Scientific); ionization: electrospray ionization with a heated electrospray interface (HESI-II) in negative polarity; spray parameters: sheath gas/aux gas/sweep gas, 30/15/0 arbitrary units; capillary temperature/heater temperature, 250 °C/250 °C; and spray voltage, -4.0 kV. The system was operated in full spectral acquisition mode within a mass range of 120–1010 m /z at ultrahigh resolving power (100,000 FWHM at 200 m/z ). Accurate mass/high-resolution (AM/HR) full scans were performed without collision energy. External mass calibration was performed for each analytical run using Pierce Negative Ion Calibration Solution (Thermo Fisher Scientific). A fluorescence nanoparticle graphene quantum dots (GQDs) labelled human membrane progesterone receptor α (hmPRα), GQD-hmPRα binding assay was performed according to the method described previously 18 , 19 . For this assay, hmPRα was first heterologously expressed in yeast ( Pichia pastoris ) using a recombinant expression system, and the solubilised hmPRα was purified through column chromatography steps. GQDs were then synthesized by pyrolysis of citric acid. The free amino groups of purified hmPRα were then covalently coupled with the free carboxylic group of GQDs to prepare GQD-labelled hmPRα. Progesterone coupled bovine serum albumin (P4-BSA) was labelled with Fluorescein isothiocynate (FITC) to prepare P4-BSA-FITC by a standard reaction method at 4 °C. Finally, for the binding assay, GQD-hmPRα and P4-BSA-FITC were mixed in a phosphate-buffered solution (pH 7.4). In the 96-well plate format, each well containing 98 µL of reaction mixture and 2 µL of ligand dissolved in ethanol, maintaining GQD-hmPRα and P4-BSA-FITC concentrations of 9 mg/ml and 14 mg/ml, respectively. Fluorescence intensity in each well was subsequently measured using a fluorescence microplate reader (Varioskan™ LUX, Thermo Scientific, Waltham, USA) with an excitation wavelength of 370 nm and an emission wavelength of 520 nm. Competitive mPRα binding activity was determined as previously described using culture cell membranes stably transfected with goldfish mPRα and the [ 3 H]-labeled maturation-inducing hormone progesterone 35 . Cultured cells were triple-washed with PBS, scraped into HEAD buffer, and sonicated for 15 s. The cell suspension was first centrifuged at 1000 × g for 7 min to eliminate nuclei and heavy mitochondrial components. The resulting supernatant was then subjected to a second centrifugation at 20,000 × g for 20 min to isolate the membrane fraction. Ligand binding to mPRα was then assessed using a previously established protocol. To measure total binding, one set of tubes contained 10 nM [ 3 H]1,2,6,7-progesterone alone, while a second set included an excess (100-fold) of cold ligand to determine non-specific binding. After a 30-min incubation with the membrane fraction at 4 °C, the binding reaction was terminated by vacuum filtration through glass microfiber filters (Whatman GF/F) pre-soaked in HAED buffer containing 2.5% Tween-80. Filters were washed three times with 5 mL of wash buffer (pH 7.4) composed of HEPES (25 mM), NaCl (10 mM), and EDTA (1 mM), maintained at 4 °C. Bound radioactivity was quantified by scintillation counting, and the displacement of [ 3 H]1,2,6,7-progesterone by unlabeled mPRα ligands represented in disintegrations per minute (DPM). Zebrafish were reared and maintained under standard laboratory conditions. The fish used in the experiments were maintained in a flow-through culture system at 28.5 °C under a 14-h light/10-h dark cycle. An in vitro and in vivo zebrafish oocyte maturation and ovulation assay was performed as described previously 15 , 17 . Four-week-old female SLc:ICR mice were used for the in vivo animal experiments. The animals were housed in a controlled environment with a 14-h light/10-h dark cycle, 22 ± 3 °C temperature, and 50–60% humidity. Conventional laboratory chow was used for feeding, with a continuous drinking water supply. The mice were reared in the laboratory environment, and 18 healthy female mice were divided into 5 groups, each containing at least 3 mice, and housed separately 5 days before the start of the experiment. Superovulation was induced with some modifications to the method applied by Kanayama et al. 36 . Pregnant mare serum gonadotropin (PMSG) and human chronic gonadotropin (HCG) at a dose of 7.5 IU were administered through intraperitoneal injection at 0 and 48 h, respectively. The treatment groups included the control group with DDW treatment, 1-CB 2-HPNA (purified fraction) at 10 mg/kg body weight, the vehicle control group with sesame oil treatment, progesterone at 2 mg/kg and mifepristone at a dose of 20 mg/kg body weight. 1-CB 2-HPNA was dissolved in DDW, whereas progesterone and mifepristone were dissolved in sesame oil. All chemicals were applied orally through an intragastric needle 6 h before HCG treatment. Finally, 20 h after HCG injection, the mice were sacrificed by cervical dislocation, and the ovaries and oviducts were collected and dried with filter paper. The oviduct was then viewed under the microscope to tear the ampulla and release the cumulus–oocyte complexes (COCs). COCs from each oviduct were then dragged into a 30 µL drop of 5 mg/mL hyaluronidase prepared in 10% fetal calf serum (FCS) to separate the ovulated oocytes. Finally, the number of ovulated oocytes was counted. The fish embryo acute toxicity (FET) test was conducted with  Danio rerio  (zebrafish) embryos following OECD Test Guideline No. 236. Fertilized eggs at 3 h post-fertilization (hpf) were exposed under static conditions to five concentrations of 1-CB 2-HPNA (0.0001, 0.001, 0.01, 0.1, and 1 mM), along with a vehicle control. A positive control using 3,4-dichloroaniline (4 mg/L) was included to verify assay sensitivity. For each treatment group, 20 viable embryos were individually placed in separate wells of a 24-well plate, each containing 2 mL of test solution. Plates were maintained at 26 ± 1 °C up to 96 hpf. Mortality and sublethal developmental endpoints (coagulation, lack of somite formation, non-detachment of the tail, absence of heartbeat, oedema, and growth delays) were assessed according to OECD criteria. The results are presented as the means ± standard deviations (SDs). Student’s t test was used to determine the statistical significance of the difference in two group. Data from different treatment groups were compared via one-way ANOVA. All the analytical data and graphs were prepared using GraphPad Prism software version 10.1.0 for Mac OS (GraphPad Software, San Diego, California, USA).

In a previous study, compounds with mPRα-binding activity from the secretion material of the marine algae Padina arborescens were fractionated into two peaks via high-performance liquid chromatography (HPLC) 22 . The compounds in both the flow-through fraction (Peak 1) and the resin-binding fraction (Peak 2) showed competitive binding activity against human mPRα and inhibitory activity on fish oocyte maturation and ovulation. In this study, we tried to purify relatively large amounts of the corresponding compounds from the extracts of the tarries of Padina . Through four-step of HPLC fractionation, two peaks of the compounds were purified from the methanol extract. In this study, the flow-through fraction from the phenyl column step was thought to be identical to Peak 1 from the secretion material in a previous study. This fraction was highly purified by using a C30 column (Supplementary Fig. S1 ). The purified fraction was subjected to 1 H-NMR, 13 C-NMR, DQF-COSY, TOCSY, HMQC and HMBC analyses (Supplementary Fig. S2 – S7 ). The compound was identified as 1-carboxybutyl 2-hydroxypentanoate (1-CB 2-HPNA) (Fig.  1 , Table 1 ). Fig. 1 Structure of 1-carboxybutyl 2-hydroxypentanoate (1-cb 2-hpna). The numbers indicate the carbon position. The one-sided arrow indicates the hmbc correlation. The bold line indicates the tocsy correlation. Table 1 Chemical shift values of 1-CB 2-HPNA. Position H ( J  = Hz) C Chemical shift values of 1-CB 2-HPNA 1 175.4 2 4.26 (m) 70.5 3 1.68, 1.82 (m) 36.6 4 1.50 (m) 18.2 5 0.95 (t, 7.4) 13.9 1’ 174.0 2’ 5.08 (m) 72.9 3’ 1.88 (m) 33.1 4’ 1.47 (m) 18.6 5’ 0.96 (t, 7.0) 13.7 Structure of 1-carboxybutyl 2-hydroxypentanoate (1-cb 2-hpna). The numbers indicate the carbon position. The one-sided arrow indicates the hmbc correlation. The bold line indicates the tocsy correlation. Chemical shift values of 1-CB 2-HPNA. The structure of 1-CB 2-HPNA was confirmed by NMR experiments ( 1 H, 13 C, DQF-COSY, TOCSY, HMQC and HMBC) via chemical shift assignments (Table 1 ). Briefly, TOCSY and DQF-COSY indicated the two proton spin systems from positions 2 to 5 and 2’ to 5’ (bold line in Fig.  1 ). The characteristic chemical shifts of positions 2 (δH 4.26, δC 70.5) and 2’ (δH 5.08, δC 72.9) indicated the presence of a hydroxy residue. The HMBC correlations (H-2 to C-1 and H-2’ to C-1’) indicated the carboxy residues in the molecule (arrows in Fig.  1 ). The HMBC correlation from H-2’ to C-1 indicated an ester bond between two units of 2-hydroxypentanoic acid. Overall, the structure of 1-CB 2-HPNA was constructed via NMR data, as shown in Fig.  1 . Since the ion peak at m/z 217.1078 [M-H] – was observed by the accurate MS, the molecular mass of 1-CB 2-HPNA was confirmed to be C 10 H 18 O 5 (theoretical value for C 10 H 17 O 5 : 217.1076 in Supplementary Fig. S8 ). The mPRα-binding activity of 1-CB 2-HPNA was subsequently evaluated using the newly established GQD-hmPRα binding assay, which allows homogeneous assay of mPR-interacting compounds (Fig.  2 A). Fig. 2 Binding of 1-CB 2-HPNA to hmPRα. ( A ) Competition of the binding of P4-BSA-FITC with GQD-hmPRα by 1-CB 2-HPNA and steroids (progesterone and estradiol-17β) was examined. ( B ) Competition for [ 3 H]-progesterone binding by progesterone or 1-CB 2-HPNA is indicated by DPM. The measurements were performed in triplicate, and the means of the data are presented with their standard deviations. Asterisks represent significant differences between ethanol alone (EtOH) and 1 μM of Progesterone or 1-CB 2-HPNA (* P  ≤ 0.05). Binding of 1-CB 2-HPNA to hmPRα. ( A ) Competition of the binding of P4-BSA-FITC with GQD-hmPRα by 1-CB 2-HPNA and steroids (progesterone and estradiol-17β) was examined. ( B ) Competition for [ 3 H]-progesterone binding by progesterone or 1-CB 2-HPNA is indicated by DPM. The measurements were performed in triplicate, and the means of the data are presented with their standard deviations. Asterisks represent significant differences between ethanol alone (EtOH) and 1 μM of Progesterone or 1-CB 2-HPNA (* P  ≤ 0.05). As shown in Fig.  2 A, compared with the positive control, progesterone decreased the fluorescence intensity of the 1-CB 2-HPNA isolated from Padina in a concentration-dependent manner. The results indicated that 1-CB 2-HPNA possessed hmPRα-interacting activity similar to that of progesterone. In contrast, no decrease in fluorescence intensity was observed for the negative control estradiol. The IC50 values for 1-CB 2-HPNA and progesterone were 1.26 and 1.90 µM, respectively. The hmPRα-interacting activity was further demonstrated by a binding assay with isotope-labeled progesterone (Fig.  2 B). The physiological activities of 1-CB 2-HPNA were evaluated by an in vitro oocyte maturation assay and an in vivo oocyte maturation and ovulation assay in zebrafish. The induction of oocyte meiotic maturation was shown to be induced by mPRα-mediated signal transduction through nongenomic action as a biological process, leading to the discovery of mPRα 4 . Org OD-02–0 is a selective agonist for mPRs and induces oocyte maturation in zebrafish. Although no agonistic activity on the induction of oocyte maturation was observed when the compounds alone were incubated with oocytes (data not shown), 1-CB 2-HPNA showed dose-dependent antagonistic activity against Org OD-02-0-induced oocyte maturation in vitro with IC50 value of 23.7 nM (Fig.  3 A). Fig. 3 Inhibitory effects of 1-CB 2-HPNA on fish oocyte maturation and ovulation. ( A ) In vitro oocyte maturation assay of 1-CB 2-HPNA. 1-CB 2-HPNA (0.001 to 1 μM) or ethanol was added to Ringer’s solution, and then, oocyte maturation was induced by 0.1 μM Org OD-02–0. After two hours of incubation, oocytes with or without germinal vesicles or with germinal vesicle breakdown (GVBD) were counted, and the percentage of GVBD was calculated. Oocytes were incubated with 0.1% ethanol (EtOH) as a negative control or with 0.1 μM Org OD-02–0 alone as a positive control (Org OD-02–0). The assay was performed in triplicate using oocytes from three fish, and the average percentage of GVBD is expressed with the standard deviation. ( B ) In vivo oocyte maturation and ovulation assay of 1-CB 2-HPNA. The antagonistic activity of 1-CB 2-HPNA against oocyte maturation and ovulation induction was analyzed by in vivo treatment in zebrafish. 1-CB 2-HPNA was added to the water at a concentration of 0.001 to 1 μM, and then, maturation and ovulation were induced by the addition of 0.1 μM Org OD-02–0. After four hours of treatment with compounds via addition to the water, the %GVBD (closed column) and %ovulation (open column) were determined by scoring the oocytes that had become transparent and formed an egg membrane by egg activation. Three fish were used for each treatment. The means and standard deviations of the data are presented. Asterisks represent significant differences between the value of percentage of GVBD or Ovulation in Org OD-02–0 alone and Org OD-02–0 with 1-CB 2-HPNA treatment (**** P  ≤ 0.00001). Inhibitory effects of 1-CB 2-HPNA on fish oocyte maturation and ovulation. ( A ) In vitro oocyte maturation assay of 1-CB 2-HPNA. 1-CB 2-HPNA (0.001 to 1 μM) or ethanol was added to Ringer’s solution, and then, oocyte maturation was induced by 0.1 μM Org OD-02–0. After two hours of incubation, oocytes with or without germinal vesicles or with germinal vesicle breakdown (GVBD) were counted, and the percentage of GVBD was calculated. Oocytes were incubated with 0.1% ethanol (EtOH) as a negative control or with 0.1 μM Org OD-02–0 alone as a positive control (Org OD-02–0). The assay was performed in triplicate using oocytes from three fish, and the average percentage of GVBD is expressed with the standard deviation. ( B ) In vivo oocyte maturation and ovulation assay of 1-CB 2-HPNA. The antagonistic activity of 1-CB 2-HPNA against oocyte maturation and ovulation induction was analyzed by in vivo treatment in zebrafish. 1-CB 2-HPNA was added to the water at a concentration of 0.001 to 1 μM, and then, maturation and ovulation were induced by the addition of 0.1 μM Org OD-02–0. After four hours of treatment with compounds via addition to the water, the %GVBD (closed column) and %ovulation (open column) were determined by scoring the oocytes that had become transparent and formed an egg membrane by egg activation. Three fish were used for each treatment. The means and standard deviations of the data are presented. Asterisks represent significant differences between the value of percentage of GVBD or Ovulation in Org OD-02–0 alone and Org OD-02–0 with 1-CB 2-HPNA treatment (**** P  ≤ 0.00001). These results suggested that 1-CB 2-HPNA acts as an antagonist through its binding activity to mPRα. The physiological activity of 1-CB 2-HPNA was further confirmed by in vivo oocyte maturation and ovulation assays in zebrafish. Again, 1-CB 2-HPNA inhibited oocyte maturation and ovulation dose-dependently (Fig.  3 B). IC50 values for oocyte maturation and ovulation in vivo assay were almost same as 24.4 nM for oocyte maturation and 14.2 nM for ovulation. Also, the IC50 values for the oocyte maturation assay in vitro and in vivo were almost the same. In addition, the physiological activity of 1-CB 2-HPNA was demonstrated in mice. The results in Fig.  4 show that the administration of 1-CB 2-HPNA inhibited ovulation. Oral administration of 1-CB 2-HPNA significantly reduced the number of ovulated eggs in the mice. These results suggest that 1-CB 2-HPNA prevents ovulation in mice via the same mechanism as it does in fish. Fig. 4 Inhibition of mouse ovulation by 1-CB 2-HPNA. 1-CB 2-HPNA was administered during superovulation induction by PMSG and HCG injection 6 h before HCG injection (42 h after PMSG injection). 1-CB 2-HPNA was dissolved in DDW and administered orally through an intragastric needle at a dose of 10 mg/kg per mouse. The control group received only water. Progesterone and mifepristone were mixed with sesame oil and administered by gavage at 2 and 20 mg/kg per mouse, respectively. The vehicle control group received sesame oil only. Finally, 68 h after PMSG injection (20 h after HCG injection), the mice were sacrificed, and the ovaries and fallopian tubes were collected. The number of ovulated oocytes isolated from the fallopian tubes was counted. Three mice were used for each experimental group. Asterisks indicate significant differences (* P  ≤ 0.05, ** P  ≤ 0.001). Inhibition of mouse ovulation by 1-CB 2-HPNA. 1-CB 2-HPNA was administered during superovulation induction by PMSG and HCG injection 6 h before HCG injection (42 h after PMSG injection). 1-CB 2-HPNA was dissolved in DDW and administered orally through an intragastric needle at a dose of 10 mg/kg per mouse. The control group received only water. Progesterone and mifepristone were mixed with sesame oil and administered by gavage at 2 and 20 mg/kg per mouse, respectively. The vehicle control group received sesame oil only. Finally, 68 h after PMSG injection (20 h after HCG injection), the mice were sacrificed, and the ovaries and fallopian tubes were collected. The number of ovulated oocytes isolated from the fallopian tubes was counted. Three mice were used for each experimental group. Asterisks indicate significant differences (* P  ≤ 0.05, ** P  ≤ 0.001).

In this study, a novel natural compound from Padina arborescence that interacts with mPRα was identified as 1-CB 2-HPNA by NMR analysis. 1-CB 2-HPNA is a novel compound that has not been previously reported. We named this new compound from Padina “Padinic acid”. In previous studies, we suggested that the major component of the compound secreted by Padina is 2-hydroxypentanoic acid (2-HPA) 23 . 1-CB 2-HPNA is the dimeric equivalent of 2-HPA. In the GQD-mPRα binding assay, 1-CB 2-HPNA showed the same affinity as progesterone (Fig.  2 ). Owing to this high affinity, 1-CB 2-HPNA inhibited fish oocyte maturation and ovulation at only a ten times higher concentration of Org OD-02–0 (Fig.  3 ). Treatment with 2-HPA resulted in the prevention of oocyte maturation and ovulation in fish. The prevention of oocyte maturation in fish is concluded to be due to the binding of 1-CB 2-HPNA to mPRα. However, we cannot exclude the possibility that the inhibition of ovulation is due to the binding of 1-CB 2-HPNA to the nuclear progesterone receptor, which induces ovulation 24 . We believe that this inhibition is due to the inhibition of oocyte maturation induction, which is the first response in the sequential induction of oocyte maturation and ovulation. Interestingly, the inhibitory effect of 1-CB 2-HPNA on ovulation was also confirmed in mice. These findings suggest that mPR is also involved in oocyte maturation and ovulation in mouse oocytes. Ovulation in mice has been reported to be induced by progesterone, but the mechanism of action remains unclear. The binding of Org OD-02-0, an agonist of mPR, and 1-CB 2-HPNA, an antagonist discovered in this study, should be verified to elucidate their mechanism of action in inducing ovulation in mice. In this study, we succeeded in identifying the first antagonist of mPRα, the first molecule discovered to mediate the nongenomic action of steroids. mPR has five subtypes, and in addition to its function in the reproductive system, which is the subject of this study, mPR has been implicated in mediating the nongenomic actions of steroids in the brain and other parts of the body as well as in regulating various biological processes. Previous reports have associated endometriosis and ovarian cancer with mPRs, which are highly expressed in these specimens 25 , 26 . Antagonists of mPRα may be useful for validating these functions and may also be candidates for new drugs. First and foremost, since it is expected to halt ovulation by inhibiting egg maturation in a dose-dependent manner in zebrafish (IC 50  = 24.4 nM), it holds potential for application as a potent contraceptive. Ovulation inhibition was also observed in mice, with effects comparable to those of the known antiprogesterone agent mifepristone (Fig.  4 ). Additionally, similar to anti-progesterone drugs, 1-CB 2-HPNA may also be usable as an anticancer agent. Anti-progesterone agents also possess anticancer effects, and several pharmaceuticals targeting breast cancer, uterine cancer, and ovarian cancer are currently in development. APR19, and EC304, targeting breast cancer, are in the preclinical stage 27 . As with these substances, 1-CB 2-HPNA may also possess anticancer activity; therefore, further investigation into its anticancer effects is necessary. Its mPR binding efficacy (IC 50  = 1.26 µM) supports limited preliminary testing in ovarian cancer cell lines such as SK-OV-3, Caov-3, IGROV-1, and OV2008 28 . Translation of these findings into i n vivo models, such as human tissue xenografts in immunosuppressed mice, would require additional validation and larger quantities (gram-scale) of compound, highlighting the need for efficient large-scale purification or chemical synthesis. A significant limitation of the current study is the small quantity of purified compound obtained from natural sources (purification yield 0.003%); thus, such a study could be conducted after the development of efficient large-scale purification methods or the establishment of chemical synthesis strategies. In particular, this compound is a nonsteroidal compound and is expected to have application as a drug without the problematic side effects of steroidal compounds. Although 1-CB 2-HPNA exibits high efficacy (in vivo IC50 = 14.2 nM), our toxicity tests using zebrafish embryos revealed no detectable toxicity even at the highest testable concentration of 1 mM (Table S1 ) 29 . Based on these results, 1-CB 2-HPNA is considered to be a substance with low toxicity, similar to 2-HPA, which we have reported on previously 23 . However, further toxicity testing is required for actual pharmaceutical applications. While cell culture-based toxicity testing methods have been used for low-dose toxicity studies, methods utilizing 3D organoids have been developed in recent years 30 , 31 . Meanwhile, toxicity assessment methods using zebrafish embryos are also gaining traction. Techniques such as high-resolution imaging of entire larvae and using AI technology to detect minute morphological abnormalities, or methods for evaluating neurotoxicity, are becoming practical 32 , 33 . It is necessary to conduct toxicity assessments by leveraging these techniques. The main challenge when considering 1-CB 2-HPNA for use in pharmaceuticals is how to produce it in large quantities. The purified 1-CB 2-HPNA obtained this time amounted to only a few dozen milligrams. Going forward, either we need to purify samples with a higher content, or we need to establish a chemical synthesis method. Although not 1-CB 2-HPNA itself, 2-hydroxycaproic acid and its polymers can be obtained from the waste liquids produced during the manufacture of alcoholic beverages (Patent Number CN111410606A). Similarly, efficient purification from these liquids and other sources might be possible. Conversely, 1-CB 2-HPNA corresponds to the dimer of the previously identified 2-HPA 23 . As chemically synthesized 2-HPA is available as a reagent, synthesizing 1-CB 2-HPNA using 2-HPA as a starting material is feasible. The chemical synthesis of 1-CB 2-HPNA is considered feasible via the dehydration synthesis of 2-HPA. To achieve this, it is first necessary to synthesize compounds in which the carboxyl group and hydroxyl group of 2-HPA are selectively protected with different protecting groups. Various protecting agents require investigation, but t-BuMe₂SiCl is considered effective for protecting the hydroxyl group, while PhCH₂OH is considered effective for protecting the carboxyl group. A potential approach involves dehydration synthesis of these hydroxyl-protected compounds, followed by protection group removal using n-Bu 4 N·F (de-silylating agent) to synthesize 1-CB 2-HPNA. Establishing this synthetic route is expected to require at least six months due to the need to optimize various conditions. Large-scale purification or chemical synthesis within a short timeframe would be desirable for actual application. The identification of novel substances from seaweed, especially water-soluble substances, is currently underway. New antagonists and agonists of mPR are likely to be discovered among these compounds. Further elucidation of novel compounds should be encouraged in the future.

Membrane progestin receptors (mPRs) are seven- or eight-transmembrane plasma membrane receptors for progesterone or its analogs. mPRs are members of a novel family of progestin and adipoQ receptors (PAQRs) consisting of 11 genes that are homologous to adipoQ receptors 1 . Among the five subtypes PAQR5 to 9, mPRα corresponds to PAQR7 2 , 3 . mPRs are conserved in a wide range of vertebrate cells, from humans to fish, and are expressed in a variety of tissues 2 , 4 . To date, mPRs have been implicated in biological regulation, including oocyte maturation in fish and amphibians and the induction of reproductive behavior in mammals 5 – 8 . mPRs have long been shown to be highly expressed in cancer cells and are thought to mediate pathways related to cancer growth and invasiveness 9 , 10 . This group of mPR molecules is known to be involved in the regulation of many cellular activities, and the search for mPR-reactive substances that respond to them is ongoing 11 , 12 . In the initial studies of mPR discovery, Org OD-02-0 was already identified as an mPR-selective agonist and has been used for research purposes 10 , 13 . We have also shown that Org OD-02-0 has the ability to induce oocyte maturation in fish 14 . The experimental systems for in vitro oocyte maturation in fish and amphibians have long been shown to be induced by the nongenomic actions of progesterone and have been used as tests for nongenomic actions, leading to the discovery of mPR 5 . This assay has been used exclusively as a method for testing mPR reactive substances 15 , 16 . We developed in vivo zebrafish oocyte maturation and ovulation induction methods as screening methods for mPR-responsive substances 17 . We have also successfully expressed and purified mPRα molecules from yeast and developed a homogeneous assay for mPR-responsive substances using graphene quantum dot (GQD)-mPRα, which is a nanoparticle-associated mPRα 18 , 19 . On the other hand, we are also trying to isolate and purify mPR-reactive natural products using these screening methods 20 . We discovered the presence of mPR-reactive substances from the marine algae Padina arborescens 21 , 22 . In this study, the compounds were purified, and the chemical structure of one of the substances was determined. The results of the physiological activity of the resulting organic compound revealed that it is an inhibitor of oocyte maturation, i.e., a novel antagonist of mPRα. mPRα-specific antagonists have never been reported before and are expected to be useful research tools and novel drug candidates in mPR research in the future.

Supplementary Information 1. Supplementary Information 2. Supplementary Information 1. Supplementary Information 2.