The CBA/J and DBA/2 mouse strains are well-established models for investigating immune-mediated RSA. In this study, female CBA/J mice were mated with male DBA/2 mice (CBA/J × DBA/2) to induce RSA-like phenotypes, as previously described (1). A total of 30 female CBA/J mice (20 ± 2 g), 10 male DBA/2 mice (22 ± 2 g), and 5 male BALB/c mice (20 ± 2 g), all aged 9 weeks, were obtained from Beijing Huafukang Biotechnology Co., Ltd. During the experimental procedures, female mice were paired with male mice at a 2:1 mating ratio. During the experimental procedures, female mice were paired with male mice in a 2:1 mating ratio. The detection of a vaginal plug was designated as embryonic day 1 of pregnancy. The pregnant females were randomly divided into three experimental groups: a normal control group, a RSA control group and an ASP intervention group. The ASP group received daily oral gavage of ASP at a dose of 400 mg/kg (32), while the control groups were administered an equivalent volume of saline. Treatments commenced on the first day of pregnancy and continued for 2 weeks. At the end of the treatment period, all mice were anesthetized and euthanized in accordance with humane protocols. All experimental procedures were conducted under the approval of the Ethics Committee of Shanghai First Maternity and Infant Hospital.

Embryos and placentas were carefully harvested from the experimental mice and immediately rinsed with ice-cold phosphate-buffered saline (PBS) to remove residual blood and debris. The cleaned samples were then subjected to rapid flash-freezing in liquid nitrogen to preserve molecular integrity and stored at −80°C until further analysis.

Frozen embryo and placenta samples, each weighing 15 milligrams, were carefully transferred into 1.5 mL Eppendorf tubes. To optimize the extraction process, two small steel beads were added to each tube, along with 0.3 mL of a methanol-to-water solution (4:1, vol/vol). Additionally, a reference solution containing 0.3 mg of L-2-chlorophenylalanine dissolved in methanol was included in each tube. The samples were subsequently incubated at −20°C for 30 min to enhance the extraction efficiency.

Following the initial storage period, the samples were subjected to ultrasonic extraction in an ice-water bath for 10 min to ensure thorough processing. They were then briefly stored at −20°C for 2 min before being ground at 60 Hz for 2 min to achieve a uniform mixture. Subsequently, the samples were centrifuged at 4°C and 13,000 rpm for 10 min, facilitating the separation of the supernatant. The collected supernatant was concentrated and dried using a freeze-drying centrifuge, yielding a final volume of 250 μL.

Each dried sample was processed with 300 μL of a methanol–water mixture (1:4, vol/vol). The mixture was vortexed for 30 s and subjected to ultrasonic extraction in an ice-water bath for 3 min. The samples were then incubated at −20°C for 2 h to ensure thorough extraction. Following incubation, the samples were centrifuged at 13,000 rpm for 10 min at 4°C. A 150 μL aliquot of the supernatant was carefully collected with a crystal syringe, passed through a 0.22 μm microfilter, and transferred to LC vials. These vials were subsequently stored at −80°C to maintain sample integrity prior to liquid chromatography-mass spectrometry (LC–MS) analysis.

To ensure the reliability and consistency of the LC–MS analysis, a pooled sample derived from equal portions of each individual sample was prepared and utilized as a quality control (QC) sample.

Metabolic profiling of the collected samples was conducted using a cutting-edge UPLC i-Series system (Waters Corporation, Milford, United States) coupled with the advanced VION IMS QTOF mass spectrometer (Waters Corporation, Milford, United States). Chromatographic separation was achieved on a high-resolution UPLC BEH C18 column (1.7 μm, 2.1 × 100 mm) in both positive and negative ionization modes. The mobile phase comprised water with 0.1% formic acid as solvent A and a 2:3 (vol/vol) acetonitrile/methanol mixture containing 0.1% formic acid as solvent B. A linear gradient elution program was employed as follows: 0 min, 1% B; 1 min, 30% B; 2.5 min, 60% B; 6.5 min, 90% B; 8.5 min, 100% B; 10.7 min, 100% B; 10.8 min, 1% B; and 13 min, 1% B. The column was maintained at a constant temperature of 45°C, with a flow rate of 0.4 mL/min. To ensure sample integrity, all samples were stored at 4°C throughout the analysis, with an injection volume precisely set to 1 μLs.

Mass spectrometry data acquisition was conducted using both full-scan mode (m/z range: 50–1,000) and MSE mode to ensure comprehensive coverage and enhanced fragmentation information. In MSE mode, alternating low- and high-energy scans were performed, enabling simultaneous acquisition of precursor and fragment ion data. The low-energy scans were conducted with a fixed collision energy of 4 eV, while high-energy scans employed a collision energy ramp ranging from 20 to 45 eV. Collision-induced dissociation was facilitated using high-purity argon gas (99.999%), with optimized instrument settings as follows: source temperature set to 115°C, desolvation gas temperature maintained at 450°C, cone voltage at 40 V, desolvation gas flow rate at 900 L/h, a scan interdelay of 0.02 s, and a scan time of 0.2 s.

To ensure data reproducibility and evaluate analytical repeatability, QC samples were systematically injected at regular intervals throughout the analysis, typically after every three sample injections. The QC samples, prepared as pooled extracts from all experimental samples, were used to monitor the relative standard deviations of both retention times and peak areas.

The specifications and details of the primary instruments utilized in this study are available in Supplementary material S1.

The raw LC–MS data were processed using Progenesis QI V2.3 software (Nonlinear Dynamics, Newcastle, United Kingdom), incorporating a comprehensive workflow that included baseline filtering, peak detection, integration, retention time correction, peak alignment, and normalization. The data processing pipeline utilized stringent parameters, including a 5% production threshold, 10 ppm product tolerance, and 5 ppm precursor tolerance, to ensure high fidelity and reproducibility. Compound identification was performed through a qualitative analysis using multiple reference databases, including the Human Metabolome Database (HMDB), LipidMaps (V2.3), Metlin, EMDB, PMDB, and a custom in-house database. Accurate mass-to-charge ratios (m/z), secondary fragment patterns, and isotopic distributions were employed as definitive criteria for compound annotation, ensuring precise and reliable metabolite identification.

The acquired data underwent rigorous preprocessing to ensure reliability and accuracy. Peaks with more than 50% missing values across groups (ion intensity = 0) were excluded. Zero values were imputed with half of the minimum detected value, and compounds were filtered based on qualitative criteria. Specifically, compounds scoring fewer than 36 points on a 60-point scale were deemed invalid and subsequently removed. Data from both positive and negative ion modes were integrated into a unified data matrix. To evaluate the overall distribution and confirm the stability of the analytical workflow, the consolidated matrix was subjected to principal component analysis (PCA) using the R programming environment.

To identify differential metabolites between experimental groups, we applied orthogonal partial least squares discriminant analysis (OPLS-DA) and partial least squares discriminant analysis (PLS-DA). Model quality was rigorously evaluated through 7-fold cross-validation and 200 response permutation tests to mitigate the risk of overfitting. The variable importance in projection (VIP) scores derived from the OPLS-DA model were utilized to quantify each variable’s contribution to group separation. Metabolites were considered differentially expressed if they met the criteria of a VIP score greater than 1.0 and a p-value less than 0.05, determined using a two-tailed Student’s t-test.

Pathway enrichment analysis of differential metabolites was conducted using their KEGG IDs, leveraging the KEGG database¹ and the analytical platform developed by Shanghai Oebiotech Co., Ltd.² Enrichment of metabolic pathways was determined using a hypergeometric test, with a significance threshold set at p ≤ 0.05. A lower p-value indicated a higher degree of significance in the differences observed across metabolic pathways. Detailed calculation formulas and methodologies are provided in Supplementary material S2.

This study was conducted with the approval of the Ethics Committee of the Obstetrics and Gynecology Hospital Affiliated to Tongji University (Ethical Approval Number: 22Y11922400). Clinical samples, including villi and decidual tissues, were collected from January 2024 to May 2024 at Shanghai First Maternity and Infant Hospital (also known as the Obstetrics and Gynecology Hospital Affiliated to Tongji University). The study population consisted of 8 patients diagnosed with RSA (RSA group) and 8 women with normal pregnancies (NC group).

The inclusion criteria for the RSA group comprised patients with a history of two or more consecutive unexplained spontaneous miscarriages occurring prior to 28 weeks of gestation. For the control group, participants were individuals undergoing elective termination of normal pregnancies, carefully matched to the RSA group based on baseline characteristics and with no prior history of spontaneous miscarriage.

Exclusion criteria encompassed any history of infections, reproductive tract abnormalities, endocrine disorders, or other identified causes of miscarriage. Baseline clinical characteristics for both groups are presented in Table 1, with additional details available in Supplementary material S3.

Total protein was extracted using RIPA lysis buffer (WB6001, Shanghai Wayo Biotechnology, Shanghai, China), and protein concentrations were quantified using the bicinchoninic acid (BCA) method (23,235, Thermo Scientific, Waltham, United States). Equal amounts of protein samples were resolved on SDS-PAGE gels and subsequently transferred onto PVDF membranes (IPVH00010, Millipore, Massachusetts, United States). Membranes were blocked with 5% non-fat milk at room temperature for 1 h, followed by overnight incubation at 4°C with primary antibodies (42,867, Cell Signaling Technology, Boston, United States). The following day, membranes were incubated with secondary antibodies for 1 h at room temperature. Immunoreactive proteins were visualized using the Tanon 5,200 imaging system (Tanon, Shanghai, China).

Grayscale intensities of protein bands were quantified using ImageJ software (NIH, Manassas, MD, United States). The relative expression of target proteins was normalized to internal controls, and mean values along with standard deviations were calculated for each group. Statistical comparisons were performed using two-tailed t-tests, with statistical significance defined as p < 0.05. All antibodies used in this experiment were obtained from the autophagy antibody kit supplied by Cell Signaling Technology.

HTR8-Svneo cells, a human chorionic trophoblast-derived cell line, were procured from the cell bank of Shanghai First Maternity and Infant Hospital. The cells were maintained in DMEM/F12 medium (C3130-0500, Biological Industries, Kibbutz Beit Haemek, Israel) supplemented with 10% fetal bovine serum and 1% penicillin–streptomycin (15140-122, Grand Island Biological Company, Montana, United States). Cultures were incubated in a humidified atmosphere of 95% air and 5% carbon dioxide at 37°C to ensure optimal growth conditions.

ASP (Yuanye, Shanghai, China) was dissolved in complete culture medium to prepare a series of concentrations: 0 μg/mL, 0.001 μg/mL, 0.01 μg/mL, 0.1 μg/mL, 1 μg/mL, and 10 μg/mL. The solutions were then sterilized by filtration through a 0.22 μm pore-sized membrane filter to ensure sterility prior to subsequent experiments.

HTR8 cells were seeded into 96-well plates at a density of 3,000 cells per well, with 100 μL of culture medium supplemented with specified concentrations of ASP. Each group included six replicates. After cell adhesion, the Cell Counting Kit-8 (CCK8, MedChemExpress, New Jersey, United States) reagent was added to the wells following the manufacturer’s protocol. A blank control, containing culture medium and CCK8 reagent without cells, was included to account for background absorbance. The optical density (OD) at 450 nm was measured using a microplate reader, with the first measurement recorded as Day 1. Subsequent measurements were performed at 24 h intervals to monitor cell proliferation dynamics.

The net OD was determined by subtracting the OD value of the blank control from that of the experimental wells. Comparative analysis of OD values across groups was conducted, and proliferation curves were generated using GraphPad Prism (version 8.0.2). Statistical significance was assessed via repeated measures analysis, followed by the least significant difference (LSD) method for post-hoc comparisons.

To evaluate cell migration, a wound healing assay was performed. Cells were seeded into 6-well plates at a density of 600,000 cells per well and cultured in serum-free medium containing varying concentrations of the tested drug. Once the cell monolayer reached approximately 90% confluence, a sterile 200-μL pipette tip was used to create a uniform, vertical scratch across the well. Detached cells and debris were carefully removed by washing the wells 2–3 times with PBS. Images of the wound area were captured using an inverted microscope (Leica, Wetzlar, Germany) at predefined time intervals. At each time point, the culture medium was replenished to maintain optimal conditions. The wound area was quantified using ImageJ software to assess the rate of wound closure over time.

To assess cell migration, 800 μL of medium containing 20% serum and the respective drug treatment was added to the lower chamber of the Transwell system, which was then placed in a 24-well plate. A total of 200 μL of cell suspension (containing 100,000 cells per well) was seeded into the upper chamber. The system was incubated at 37°C in a humidified incubator with 5% CO₂ for 16 h. Following incubation, cells that had migrated to the lower surface of the membrane were fixed with 4% paraformaldehyde at room temperature for 20 min. The fixed cells were stained with 0.1% crystal violet solution for 30 min. Non-migrated cells on the upper surface of the membrane were carefully removed using a cotton swab. The membranes were then rinsed with PBS to eliminate excess stain. Migrated cells were visualized and imaged under an inverted microscope for quantitative analysis.

In contrast to the migration assay, the invasion assay incorporates an additional step to assess cell invasive capabilities. The upper chamber is pre-coated with 100 μL of medium containing 10% Corning matrigel matrix (Corning, New York, United States) to mimic the extracellular matrix. The chamber is incubated at 37°C for 1 h to allow the Matrigel to solidify, after which the supernatant is carefully removed. The subsequent procedures, including cell seeding and incubation, follow the same protocol as described for the migration assay.

Image analysis was conducted using ImageJ software, while data visualization and statistical evaluations were performed with GraphPad Prism (version 8.0.2). Results are presented as the mean ± standard error of the mean. Statistical comparisons between groups were performed using one-way analysis of variance. Post hoc analyses were carried out using either the LSD or Bonferroni multiple comparison tests to assess the significance of intergroup differences. Statistical significance was defined as follows: p < 0.05 (*), p < 0.01 (**), or p < 0.001 (***).

Cells were harvested via centrifugation and promptly fixed in a 2.5% glutaraldehyde solution at 4°C for a minimum of 3 h to preserve cellular structures. Following fixation, samples were washed three times with 0.1 M phosphate buffer and subsequently post-fixed in 1% osmium tetroxide at 4°C for 3 h. The specimens were then subjected to three additional washes with phosphate buffer, sequentially dehydrated in graded ethanol, and embedded in Epon 812 resin to ensure optimal preservation and sectioning quality.

Ultrathin sections, approximately 70 nm in thickness, were prepared using an ultramicrotome (Leica UC6) and carefully mounted onto copper grids coated with formvar support films. The sections were stained with uranyl acetate for 30 min to enhance contrast, followed by counterstaining with lead citrate for 15 min. Finally, the stained sections were visualized and imaged using a transmission electron microscope (Thermo Fisher Talos 120) operated at 120 kV.

The diagnostic criteria for RSA were defined according to the ESHRE guidelines (33, 34). The clinical characteristics of participants in the RSA group and the NC group are summarized in Table 1 and Supplementary material S3. No significant differences were observed between the two groups in terms of age (30–34 years), body mass index (BMI, 20–24), or gestational age (7–10 weeks) (p > 0.05).

To investigate autophagy activity at the maternal-fetal interface, we examined decidual tissues from RSA patients and compared them to those of the NC group. As illustrated in Figure 1, autophagy flux was assessed across 8 samples from the NC group and 8 samples from the RSA group. Western blot analysis revealed a significant reduction in the expression levels of autophagy-related proteins, including ATG5, ATG7, and ATG16L, in the RSA group. Notably, Beclin 1 levels were significantly decreased (p-value <0.05). Although the expression of several other proteins implicated in the autophagy pathway did not show significant differences, the overall autophagy levels in the RSA group displayed a clear downward trend.

Previous studies have demonstrated that ASP promotes autophagy activation (35) and improves outcomes in RSA animal models (36). Furthermore, ASP has been reported to confer protective effects during pregnancy (32). Based on these findings, we hypothesize that ASP may mitigate RSA by activating protective autophagy pathways. Data presented in Supplementary material S4 illustrate the miscarriage status of mice in the RSA model. WB analysis revealed that the expression levels of autophagy-related proteins, including ATG7, ATG16L, and Beclin 1, were significantly elevated in the ASP-treated group compared to the untreated RSA group (Figures 1C,D). Additionally, our previous metabolomic analysis highlighted enrichment of differential metabolites in the autophagy pathway, with pathway activity upregulated in the ASP-treated group relative to controls (Supplementary material S5).

To investigate the metabolic alterations induced by ASP in the context of RSA, a metabolomic analysis was conducted comparing samples from the ASP-treated and control groups (Figure 2A). An OPLS-DA model revealed a clear and optimized class separation, demonstrating robust model fitting and effectively capturing the metabolic changes induced by ASP exposure (Figure 2B). Among the identified metabolites, 55 were significantly downregulated, and 42 were upregulated in the ASP group compared to the control group (Figure 2C; Supplementary materials S6, S7). Supporting our hypothesis, phosphatidylethanolamine (PE) was prominently altered between the two groups. The differential metabolites identified belong to several chemical classes, including benzene and substituted derivatives, carboxylic acids and derivatives, and fatty acyls, etc. (Supplementary materials S8, S9). Pathway enrichment analysis using the KEGG database highlighted significant enrichment of these metabolites in pathways such as glycolysis/gluconeogenesis, glycerolipid metabolism, glycine, serine, and threonine metabolism, nicotinate and nicotinamide metabolism, glyoxylate and dicarboxylate metabolism, Fc gamma R-mediated phagocytosis, and the Apelin signaling pathway (Figure 2D; Supplementary material S10). Subsequently, autophagosomes were observed by TEM. TEM further provided direct evidence of autophagic activity. Autophagosomes were visualized in the decidual tissues of the normal mouse model (Figure 2E), the RSA mouse model (Figure 2F), and the ASP-treated RSA mouse model (Figure 2G).

The impact of ASP on the proliferation of HTR8 cells was assessed using the CCK-8 assay, revealing that ASP significantly promoted cell proliferation in a dose-dependent manner (Figure 3A). Consistently, transwell migration and scratch wound healing assays demonstrated a marked enhancement in the migratory capacity of HTR8 cells upon ASP treatment (Figures 3B,C,F,G). Additionally, the transwell invasion assay further confirmed that ASP significantly facilitated the invasive ability of HTR8 cells (Figures 3D,E). Collectively, these findings indicate that ASP serves as a potent enhancer of trophoblast cell proliferation, migration, and invasion, underscoring its potential role in promoting trophoblast function in a dose-dependent manner.