Chlorogenic Acid Ameliorates Chronic Unpredictable Stress-Induced Diminished Ovarian Reserve Through Ovarian Renin-Angiotensin System.

The research was in compliance with the Helsinki Declaration and approved by the Ethics Committee of the First Affiliated Hospital of Soochow University. All animal experiments were approved by Soochow University Institutional Animal Care and Use Committee (SUDA20240911A25).

To evaluate the effects of CUS‐induced DOR model in mice, we assessed behavioral changes through SCT and TST, measured body weight, and detected serum CORT and sex hormone levels to evaluate the stress level and HPO endocrine function. First, the body weight change curve was illustrated in Figure  S3A , the average body weight of mice under low, medium, and high‐frequency stimulation was lower than that of the control group after 8 weeks of CUS intervention, with the most significant difference in the high‐frequency stimulation group ( p  < 0.0001). Second, we validated the effects of CUS on the cognitive function and emotions of mice. SCT results showed that the percentage of sucrose consumption decreased in Group M (Figure  S3B , p  < 0.05) and Group H (Figure  S3B , p  < 0.01), indicating depressive emotions such as anhedonia. Compared with Group C, the immobility time in TST for Group M ( p  < 0.05) and Group H ( p  < 0.001) was prolonged (Figure  S3C ), manifesting as a state of despair. Both behavioral experiments indicated that CUS intervention resulted in depressive‐like behavior in mice. In addition to inhibiting the growth potential of mice and interfering with emotional behavior changes, CUS‐induced changes in endocrine function are also an important aspect of CUS outcomes. We found that compared with Group C, the serum CORT levels in Group M and Group H ( p  < 0.0001 and p  < 0.0001) mice significantly increased (Figure  S3D ), indicating that medium and high‐frequency CUS stimulation would cause individuals to remain in a continuous stress state. Although the short‐term elevation of serum CORT levels can quickly mobilize the body's defense mechanisms to better cope with sudden external situations, producing beneficial short‐term effects, long‐term exposure to high concentrations of CORT may impair the body's acutely protective physiological systems [ 46 ]. In terms of ovarian endocrine function, the serum AMH (Figure  S3E ) and E 2 (Figure  S3G ) levels in Group H were significantly lower than those in Group C ( p  < 0.0001 and p  < 0.0001), while the FSH (Figure  S3F ) levels were significantly higher than those in Group C ( p  < 0.001). Notably, AMH seems to be more susceptible to the effects of CUS and shows a certain frequency effect, leading to decreased expression under low, medium, and high‐frequency CUS (Figure  S3E ). These results suggest that CUS has significant effects on the growth potential and emotional behavior of mice, and it was found to induce DOR in mice at the hormone level. To investigate whether CUS affects the estrous cycle in mice, we used vaginal smears and observed different cell types to determine the specific stages of the estrous cycle. As shown in Figure  S3H , H&E staining demonstrated the characteristic cell types at different stages, including proestrus (P), estrus (E), metestrus (M), and diestrus (D). Compared with Group C, the estrous cycle in Group H was completely disrupted and mostly stayed in diestrus (Figure  S3I ), indicating that CUS could affect ovarian endocrine function and cause estrous cycle disorder in mice. Under normal feeding conditions, the ratio of ovarian wet weight to body weight in mice is relatively constant. By measuring the ovarian organ index, we found that compared with Group C, the ovarian index decreased after low ( p  < 0.01), medium ( p  < 0.001), and high‐frequency ( p  < 0.0001) stimulation (Figure  S3J ), indicating that CUS could cause ovarian dysplasia or atrophy and show a certain frequency effect. Histologically, the ovarian tissue in Group H showed overall structural atrophy and thinning of the cortex (Figure  S3K ). Compared with Group C, the number of functional follicles in Group H decreased (Figure  S3L ), including primordial follicles ( p  < 0.01), primary follicles ( p  < 0.05), secondary follicles ( p  < 0.01), and mature follicles ( p  < 0.05). Notably, the number of atretic follicles and corpora lutea increased in Group H ( p  < 0.01 and p  < 0.01). The above results indicate at the histological level that CUS can lead to DOR, with a more pronounced loss of secondary and mature follicles, consistent with the decrease in AMH levels mentioned above. Additionally, the reduction in the number of primordial follicles and the increase in corpora lutea in Group H indicate that CUS can accelerate follicle depletion, causing DOR. In summary, 8 weeks of continuous CUS disrupts the normal morphology of ovarian tissue and causes endocrine function damage in mice. To investigate the effect of CUS on the quality and quantity of oocytes in mice, we collected MII oocytes from each group through superovulation. Figure  S4A,B illustrated that the number of oocytes obtained in Group M and H was lower than that in Group C ( p  < 0.05 and p  < 0.01). This indicates that CUS‐induced high concentrations of glucocorticoids severely impair ovarian reserve, leading to reduced superovulation efficiency. Tissue damage caused by glucocorticoids is associated with oxidative stress [ 47 ]. To assess oxidative damage in oocytes of each group, we first used a fluorescent probe to detect ROS levels in oocytes. The results showed that ROS levels in oocytes of Group M and H were significantly higher compared to Group C, indicating that CUS increases oxidative stress levels in the ovaries, particularly in oocytes (Figure  S4C,D , p  < 0.01 and p  < 0.001). Mitochondria are considered the primary source of ROS in cells, and their proper function is crucial for cellular energy activities. Therefore, we measured mitochondrial membrane potential levels in oocytes of each group. In Figure  S4E,F , compared to Group C, mitochondrial membrane potential in the oocytes of Group L, M, and H decreased, with marked mitochondrial depolarization observed ( p  < 0.05, p  < 0.01, and p  < 0.001). This suggests that the high concentration of glucocorticoids induced by CUS may increase intracellular ROS levels by reducing mitochondrial membrane potential in oocytes, thereby elevating oxidative stress levels in oocytes, with a certain frequency effect. Similarly, to investigate whether chronic stress interferes with GC function, we used Dex to induce KGN cells and establish an in vitro chronic stress model. In Figure  S4G,H , we found that after adding Dex, ROS accumulation levels in KGN cells increased ( p  < 0.01). Mitochondrial membrane potential levels showed similar results. Compared to the Control group, mitochondrial membrane potential levels in the Dex group decreased (Figure  S4I,J , p  < 0.05). These results suggest that the high levels of glucocorticoids caused by chronic stress may lead to DOR by increasing oxidative stress levels in oocytes and GCs. To further explore the mechanism by which OVRAS is involved in the DOR process induced by high concentrations of glucocorticoids, we analyzed the expression changes of key components of OVRAS at both mRNA and protein levels using both the in vivo model of DOR mice induced by CUS and the in vitro model of KGN cells induced by Dex. First, in the ACE‐AngII‐AT1R pathway, at the mRNA level, as was known in Figure  S5A , the expression levels of Ace and Agt in Group H were higher than those in Group C ( p  < 0.01 and p  < 0.01), and the expression levels of both Agtr1 receptor subtypes a and b were also increased ( p  < 0.05 and p  < 0.01). Regarding protein expression (Figure  S5C,G ), compared with Group C, the expression levels of ACE and AGT in Group H were increased ( p  < 0.05 and p  < 0.01). AGTR1 protein expression increased with the frequency of CUS, showing a significant frequency effect ( p  < 0.01). Additionally, ovarian tissue homogenate results indicated that the local expression level of AngII increased with stimulation frequency (Figure  S5E ), especially in Group M ( p  < 0.001) and Group H ( p  < 0.01). We also examined changes in serum AngII levels, which were similar to the results from the ovarian tissue homogenates. As the CUS stimulation frequency increased, serum AngII levels also increased (Figure  S5H ), with more significant increases observed in Group M and H ( p  < 0.0001 and p  < 0.0001). These results indicate that CUS induced overactivation of the ACE‐AngII‐AT1R axis at both the transcription and translation levels. Notably, compared with Group C, the expression levels of Agtr2 in Group L increased at both the mRNA (Figure  S5B ) and protein (Figure  S5D ) levels ( p  < 0.05 and p  < 0.05). Similarly, we examined the expression levels of key components of the ACE2‐Ang(1‐7)‐MasR pathway. At the mRNA level (Figure  S5B ), the expression level of Ace2 in the L group was significantly increased compared with Group C ( p  < 0.01), and similar results were observed at the protein level (Figure  S5D , p  < 0.05). However, there were no changes in MasR levels at either the mRNA (Figure  S5B ) or protein (Figure  S5D ) level between groups. Except for the increase in Ang(1‐7) expression in Group L ( p  < 0.05), there were no significant changes in the local ovarian expression levels of Ang(1‐7) between groups (Figure  S5F ), and serum Ang(1‐7) levels also remained unchanged. In conclusion, CUS induced changes in the expression of key components of OVRAS in DOR mice, especially in the ACE‐AngII‐AT1R pathway. In the in vitro cell model, consistent with the in vivo animal model results, we observed that in KGN cells treated with Dex, ACE , AGT , and AGTR1 expression levels increased at the RNA level (Figure  S5J ) ( p  < 0.001, p  <0.01, and p  < 0.001), with similar results at the protein level (Figure  S5L,N ). For the ACE2‐Ang(1‐7)‐MasR pathway, we found that high concentrations of glucocorticoids did not induce significant changes in expression levels at either the RNA level (Figure  S5K ) or the protein level (Figure  S5M,N ). These results indicate that high concentrations of glucocorticoids could induce activation of the ACE‐AngII‐AT1R pathway in the granulocyte RAS. Next, we supplemented CGA by gavage on the basis of CUS intervention to explore whether CGA could play a therapeutic role in CUS‐induced DOR. First, by observing the body weight change curve (Figure  1A ), the trend in Group H was consistent with previous studies. After 1 week of CGA supplementation, the average body weight of Group A began to be slightly higher than that of Group H, by the end of the 8th week ( p  < 0.05). By comparing the ovarian organ index, it was found that the ovarian index of Group A ( p  < 0.01) increased compared to Group H, indicating that CGA can improve CUS‐induced ovarian atrophy and dysplasia (Figure  1B ). Next, focusing on endocrine function, compared with Group H, the CORT level in Group A significantly decreased (Figure  1C , p  < 0.01), indicating that the chronic stress state induced by CUS was alleviated. However, FSH (Figure  1E ) and E 2 (Figure  1F ) were not significantly improved after CGA supplementation, suggesting that CGA seems to be more beneficial to the recovery of ovarian reserve. Meanwhile, the secretion level of AMH significantly increased (Figure  1D , p  < 0.001). Further studies showed that in terms of the estrous cycle, compared with Group C, the estrous days of Group H decreased ( p  < 0.05), and the diestrus days increased ( p  < 0.01), which is similar to previous study results. However, CGA reduced the diestrus days ( p  < 0.01), indicating that CGA supplementation improved the ovarian endocrine function of the mice. Ovarian histology and follicle counts at all levels indicated that after CGA treatment (Figure  1K,H ), the numbers of primordial follicles ( p  < 0.05), secondary follicles ( p  < 0.05), and mature follicles ( p  < 0.05) in Group A increased compared with Group H, and the number of atretic follicles correspondingly decreased ( p  < 0.05), indicating that CGA could improve ovarian morphology and follicle development to some extent. In addition, since GCs apoptosis often leads to follicle atrophy and ovarian function decline [ 48 ], we performed apoptosis staining on ovarian tissues. The proportion of positive cells in Group H mice significantly increased ( p  < 0.001). Notably, Figure  1J showed that apoptotic cells were mostly concentrated in the GCs of the follicular hilus. After CGA treatment, the proportion of apoptotic GCs in the ovary decreased (Figure  1I , p  < 0.05). GCs play an important role in maintaining ovarian function, and the development and maturation of oocytes are affected by the function of GCs. CGA could reduce the apoptosis level of GCs induced by CUS, thereby improving the development and maturation of oocytes. In summary, CGA provides a protective effect against ovarian damage caused by CUS, specifically by partially restoring growth potential in mice, reducing GCs apoptosis levels, and improving ovarian morphology and endocrine function. CGA ameliorated symptoms of DOR induced by CUS. (A) Dynamics of mouse weight variation during modeling ( n = 15), *Group C versus Group H at every week and # Group H versus Group A at the end of the 8th week. (B) The ovarian index was measured at the end of intervention period ( n = 6). Serum hormone indicators of CORT (C), AMH (D), FSH (E), and E 2 (F), which were related to chronic stress and ovarian reserve ( n = 6). (G) Cumulative days of each estrous cycle in different groups ( n = 3). (H) The number of follicles at various stages were counted via H&E staining ( n = 3). (I) and (J) Apoptosis was evaluated by Tunel staining (scale bar: 200 µm and scale bar: 25 µm) and apoptotic rate was calculated for percentage of Tunel‐positive cells in each group of mouse ovarian tissue ( n = 3). (K) H&E‐stained sections of mice ovaries (scale bar: 200 µm) and magnified view of the selected region (scale bar: 50 µm). The follicles at all levels: primordial follicle→1, primary follicle→2, secondary follicle→3, mature follicle→4, atretic follicle→5. Results are expressed as the mean ± SD and one way ANOVA was used for between‐group comparisons, n represents the number of biological replicates. * p  < 0.05, ** p  < 0.01, *** p  < 0.001, **** p  < 0.0001, ns: not significant. Chronic stress affects oocyte quality in mice [ 49 ]. To investigate whether CGA has a therapeutic effect, we obtained MII oocytes from mice in each group through superovulation. First, in terms of the number of oocytes retrieved, as was illustrated in Figure  2A,C , we found that after supplementing with CGA, the number of oocytes retrieved was significantly increaed compared to Group H ( p  < 0.01), indicating that CGA could partially restore ovarian reserve. Fertilization rate can be used as one index to assess oocyte quality. Through in vitro fertilization of MII oocytes (Figure  2B,D ), we found that there was no significant difference between Group H and Group C. Furthermore, the fertilization rate in Group A did not improve compared to Group H ( p  > 0.05), indicating that CUS does not significantly affect the fertilization rate of oocytes. Tissue damage induced by chronic stress is closely associated with oxidative stress [ 25 ]. To assess the oxidative damage to oocytes in each group, we used fluorescent probes to detect ROS levels within the oocytes (Figure  2E,G ). The results showed that compared to Group C, exogenous supplementation with CGA slightly alleviated the oxidative stress status in oocytes ( p  < 0.05). Mitochondria are considered a major source of ROS within cells. Therefore, we assessed the mitochondrial membrane potential levels in oocytes from each group of mice (Figure  2H,F ). As was known in Figure  2H,F , compared to Group C, after CGA treatment, mitochondrial depolarization in oocytes was reduced, and the membrane potential levels increased ( p < 0.05), indicating that CGA can reduce intracellular ROS levels by increasing mitochondrial membrane potential in oocytes, thereby alleviating oxidative stress levels within the oocytes. These results indicate that CGA could restore ovarian function impaired by CUS‐induced reduction in ovarian reserve by lowering oxidative stress levels in oocytes. CGA increased the number of mature oocytes and decreased oxidative stress levels in oocytes caused by CUS. (A) and (C) Brightfield images showing oocytes of each group and counts of oocytes ( n = 3). Scale bar: 100 µm. (B) Brightfield images showing pictures of fertilized and unfertilized oocytes. Scale bar: 100 µm. (D) Oocytes fertilization rates in different groups after CUS and CGA interventions ( n = 3). (E) and (G) Immunofluorescence demonstrated the ROS levels and quantification in oocytes at each group by DCFH‐DA probe ( n = 5). Scale bar: 100 µm. (F) Mitochondrial membrane potential in oocytes analyzed by the JC‐1 aggregate/monomer fluorescence ratio ( n = 5). (H) JC‐1 staining of oocyte mitochondrial membrane potential ( n = 5). Scale bar: 100 µm. Results are expressed as the mean ± SD and one way ANOVA was used for between‐group comparisons, n represents the number of biological replicates. * p  < 0.05, ** p  < 0.01, *** p  < 0.001, **** p  < 0.0001, ns: not significant. To determine whether CGA improves DOR by reducing the activity of the ACE‐AngII‐AT1R pathway and thereby decreasing oxidative stress and apoptosis levels in ovary, we assessed the molecular expression of key components in the OVRAS pathway in mouse ovaries, as well as oxidative stress and apoptosis markers. First, after CGA treatment, the molecules that were upregulated by CUS intervention showed a certain degree of downregulation at the RNA level, specifically Ace ( p  < 0.05), Agt ( p  < 0.05), and Agtr1a ( p  < 0.01) expression levels decreased (Figure  3A ). Similar phenomena were observed at the protein level (Figure  3D,G ), and the ovarian tissue homogenate results also indicated a decrease in local ovarian AngII levels after CGA addition (Figure  3I , p  < 0.05), indicating that CGA supplementation reduces the activity of the ACE‐AngII‐AT1R pathway. Similarly, we assessed the molecular expression of key components in the ACE2‐Ang(1‐7)‐MasR pathway. Consistent with previous in vivo experiments, there were no significant differences in Ace2 and MasR expression levels at either the mRNA (Figure  3B ) or protein level (Figure  3E,G ). In Figure  3J , ovarian tissue homogenate results also showed no significant changes in Ang(1‐7). CGA improved DOR by reducing oxidative stress and apoptosis levels in mouse ovaries through its effects on the ACE‐AngII‐AT1R axis. (A) and (B) The qRT‐PCR analysis of the OVRAS components Ace , Agt , Agtr1a , Agtr1b , Ace2 , Agtr2 , and MasR ( n = 3). (C) Expression levels of oxidative stress and apoptosis molecular markers ( Bax , Bcl‐2 , Sod2 , and Gpx4 ) in the ovaries were detected by qRT‐PCR ( n = 3). (D–F) Quantification of ACE, AGT, AGTR1, ACE2, AGTR2, MasR, BAX, Bcl‐2, SOD2, and GPX4 protein expression. (G) and (H) Photographs of western blot showing the protein level of ACE, ACE2, AGT, AGTR1, AGTR2, MasR, BAX, Bcl‐2, GPX4, and SOD2. (I) and (J) Analysis of AngII and Ang(1‐7) in mouse ovary tissue homogenates collected from each group ( n = 3). CAT (K), GSH (L), and MDA (M) levels as oxidative stress indicators were determined in mouse ovary tissue homogenates ( n = 6). Results are expressed as the mean ± SD and one way ANOVA was used for between‐group comparisons, n represents the number of biological replicates. * p  < 0.05, ** p  < 0.01, *** p  < 0.001, ns: not significant. Chronic stress can induce tissue damage through oxidative stress and apoptosis [ 50 ]. To assess oxidative stress and apoptosis levels in mouse ovaries, we measured the expression levels of Sod2 , Gpx4 , Bax , and Bcl‐2 , as well as the activities of GSH and CAT, and the MDA levels, which are widely accepted biomarkers of oxidative damage and apoptosis [ 51 , 52 ]. First, at the oxidative stress level, we observed an increase in MDA levels in Group H (Figure  3M  , p  < 0.001), along with decreased CAT and GSH activities (Figure  3K , p  < 0.01 and Figure  3L , p  < 0.01). CGA reversed the increase in oxidative stress induced by CUS, with a decrease in MDA levels in Group A compared to Group H ( p  < 0.05) and a recovery in GSH activity ( p  < 0.05). There was a similar trend for Sod2 and Gpx4 expression. At the mRNA level, CGA supplementation led to a recovery of Sod2 and Gpx4 expression levels that were reduced by CUS intervention (Figure  3C , p  < 0.05 and p  < 0.05). At the protein level, we observed an increase in SOD2 levels after CGA treatment (Figure  3F , p  < 0.05). We also assessed the expression of apoptosis‐related molecules. At the RNA level, compared to Group C, the pro‐apoptotic gene Bax expression was increased in Group H (Figure  3C , p  < 0.01), while the expression of the anti‐apoptotic gene Bcl‐2 was decreased (Figure  3C , p  < 0.01). At the protein level, the expression trends were similar, with increased BAX levels (Figure  3F , p  < 0.001) and decreased Bcl‐2 levels (Figure  3F , p  < 0.05). This indicates that ovarian apoptosis levels increased under CUS intervention. After CGA intervention, we observed a decrease in Bax expression at both the RNA and protein levels (Figure  3C , p  < 0.05 and Figure  3F , p  < 0.05). Previous research has shown that the ratio of Bax/Bcl‐2 can be a key determinant of cell survival or death when apoptosis is triggered [ 53 ]. In our experiment, although Bcl‐2 levels did not show significant recovery after CGA supplementation at either RNA or protein levels (Figure  3C,F ), it was noteworthy that the Bax/Bcl‐2 ratio decreased after CGA supplementation at both RNA and protein levels (Figure  3C , p  < 0.01 and Figure  3F , p  < 0.05). This indicates that CGA could reduce the overall level of ovarian apoptosis to some extent. These results indicate that CGA reduced the activity of the ACE‐AngII‐AT1R pathway, improving ovarian damage caused by increased oxidative stress and apoptosis levels induced by CUS. To investigate whether chronic stress interferes with GC function through the OVRAS pathway, we used Dex to induce KGN cells and established an in vitro chronic stress model to assess changes in the expression of key RAS components, oxidative stress and apoptosis levels in KGN cells after Dex treatment and CGA intervention. qRT‐PCR analysis indicated that compared to the Control group, the expression levels of ACE ( p  < 0.001), AGT ( p  < 0.001), and AGTR1 ( p  < 0.01) were increased under Dex stimulation (Figure  4A ). Consistent with the mRNA levels, as shown in Figure  4D,G , the protein levels of ACE ( p  < 0.01), AGT ( p  < 0.001), and AGTR1 ( p  < 0.05) were elevated in KGN cells after Dex stimulation. Following CGA treatment, the increased expression of molecules under Dex stimulation was partially reversed at the RNA level (Figure  4A ), as evidenced by decreased levels of ACE ( p  < 0.05), AGT ( p  < 0.05), and AGTR1 ( p  < 0.05). Similar reductions in protein expression levels of ACE ( p  < 0.05), AGT ( p  < 0.01), and AGTR1 ( p  < 0.05) were also observed (Figure  4D,G ). Additionally, changes in the expression levels of BAX , Bcl‐2 , GPX4 , and SOD2 were assessed to reflect the oxidative stress and apoptosis levels in each group. We observed that with the addition of CGA, the expression of BAX decreased (Figure  4C,F , p  < 0.05 and p  < 0.05), and the expression of Bcl‐2 increased (Figure  4F , p  < 0.05). The expression level changes of GPX4 and SOD2 , which reflect oxidative stress levels, showed a similar trend. Specifically, after adding CGA, the oxidative stress level in KGN cells was partially restored at both RNA and protein levels (Figure  4C,F , p  < 0.05, Figure  4C,F , p  < 0.05). The results indicate that the activity of the ACE‐AngII‐AT1R pathway, which is overactivated by Dex, is mitigated by CGA. We then assessed changes in the ACE2‐Ang(1‐7)‐MasR pathway using the same method. Similar to the previous results in mouse ovaries, no significant changes were observed at the RNA and protein levels. Moreover, unlike the in vivo experiments, no significant differences in the expression levels of ACE2 and AGTR2 were observed in the in vitro studies. In conclusion, the effects caused by Dex may stem from the excessive activation of the ACE‐AngII‐AT1R pathway in KGN cells, disrupting the equilibrium with the ACE2‐Ang(1‐7)‐MasR axis. CGA decreased activity of ACE‐AngII‐AT1R axis as well as reduced oxidative stress and apoptosis levels of Dex‐induced KGN cells. (A) and (B) The qRT‐PCR analysis of OVRAS components ACE , AGT , AGTR1 , ACE2 , AGTR2 , and MasR mRNA expression level ( n = 3). (C) Expression levels of oxidative stress and apoptosis molecular markers ( BAX , Bcl‐2 , SOD2 , and GPX4 ) detected by qRT‐PCR ( n = 3). (G) and (H) The OVRAS, oxidative stress and apoptosis‐related protein expression in PVDF membrane. (D) and (E) Quantification of ACE, AGT, AGTR1, ACE2, AGTR2, and MasR protein expression ( n = 3). (H) Quantification of BAX, Bcl‐2, SOD2, and GPX4 protein expression ( n = 3). Results are expressed as the mean ± SD and one way ANOVA was used for between‐group comparisons, n represents the number of biological replicates. * p  < 0.05, ** p  < 0.01, *** p  < 0.001, ns: not significant. To further investigate whether CGA reduces oxidative stress and apoptosis levels in KGN cells through the ACE‐AngII‐AT1R pathway, we co‐treated KGN cells with the AGTR1 agonist AngII and the inhibitor CS along with CGA. Initially, we assessed ROS accumulation in cells across different groups, as illustrated in Figure  5A . Compared to the Control group, ROS levels were significantly elevated in the Dex group (Figure  5B , p  < 0.001), but CGA treatment reduced ROS levels (Figure  5B , p  < 0.01). Additionally, the addition of the AGTR1 agonist AngII further elevated ROS levels compared to the Dex+CGA group (Figure  5B , p  < 0.001). Mitochondrial membrane potential levels mirrored the ROS results (Figure  5C ), with a notable decrease in potential after AngII addition compared to the Dex+CGA group (Figure  5D , p  < 0.001). The AGTR1 inhibitor CS was able to reverse this decrease and lower oxidative stress levels (Figure  5D , p  < 0.05). Additionally, we observed that the expression levels of intracellular antioxidants such as GPX4 and SOD2 decreased after adding Dex (Figure  5J , p  < 0.001 and Figure  5K , p  < 0.001), indicating that Dex induces oxidative damage in KGN cells. After CGA supplementation, the expression levels of GPX4 and SOD2 showed some recovery (Figure  5J , p  < 0.05 and Figure  5K , p  < 0.05). In addition, adding CS did not result in statistically significant changes in GPX4 and SOD2 levels (Figure  5J,K ), whereas adding AngII significantly reduced GPX4 and SOD2 levels (Figure  5J , p  < 0.01 and Figure  5K , p  < 0.01). These results suggest that excessive activation of AGTR1 led to oxidative damage in cells, and CGA could improve oxidative damage in KGN cells by reducing the activity of the ACE‐AngII‐AT1R pathway. CGA decreased oxidative stress and apoptosis levels caused by ACE‐AngII‐AT1R axis in KGN cells. (A) and (B) Detection of intracellular ROS by DCFH‐DA probe and quantitative analysis results of the ROS in different groups ( n = 3), * vs. Group Dex and + vs. Group Dex+CGA. Scale bar: 100 µm. (C) and (D) Measurement and the analysis of mitochondrial membrane potential levels in each group ( n = 3), * vs. Group Dex and + vs. Group Dex+CGA. Scale bar: 100 µm. (E) Apoptosis levels determined by Annexin V‐FITC assay labeling in KGN cells. Scale bar: 100 µm. The oxidative stress and apoptosis molecular markers protein levels were measured by western blot (F) and the analysis of BAX (G), Bcl‐2 (H), BAX/Bcl‐2 (I), GPX4 (J), and SOD2 (K) from different groups ( n = 3), * vs. Group Dex and + vs. Group Dex+CGA. Results are expressed as the mean ± SD and one way ANOVA was used for between‐group comparisons, n represents the number of biological replicates. * p  < 0.05, ** p  < 0.01, *** p  < 0.001, + p  < 0.05, ++ p  < 0.01, +++ p  < 0.001. To investigate whether CGA reduces apoptosis damage in Dex‐induced KGN cells through the ACE‐AngII‐AT1R pathway, we used the FITC‐labeled Annexin V fluorescent probe to detect apoptosis levels in cells. In Figure  5E , the results indicated that compared to the Control group, the number of early apoptotic cells (green fluorescence) increased in the Dex group. After adding AngII, the number of late apoptotic/necrotic cells (red fluorescence) increased, suggesting that excessive activation of the ACE‐AngII‐AT1R axis in KGN cells promoted apoptosis. Supplementation with CGA and CS reversed this effect, evidenced by a reduction in late apoptotic/necrotic cells. Meanwhile, we measured the expression levels of BAX and Bcl‐2. After adding Dex, the expression level of Bcl‐2 decreased (Figure  5H , p  < 0.01) while the expression level of BAX increased (Figure  5G , p  < 0.01). Supplementation with CGA reversed the expression levels of BAX, as evidenced by a decrease in BAX expression (Figure  5G , p  < 0.05). Building on this, we added AngII and CS to investigate whether CGA further affects apoptosis levels in KGN cells through the ACE‐AngII‐AT1R pathway. After adding AngII, BAX expression significantly increased (Figure  5G , p  < 0.01) while Bcl‐2 expression significantly decreased (Figure  5H , p  < 0.001). After adding CS, there were no significant changes in the expression of individual genes such as BAX or Bcl‐2, but CS reduced the BAX/Bcl‐2 ratio (Figure  5I , p  < 0.05). These results indicate that Dex‐induced apoptosis levels in KGN cells increase, and CGA could reduce apoptosis levels in KGN cells through the ACE‐AngII‐AT1R pathway. In summary, excessive activation of the ACE‐AngII‐AT1R axis in KGN cells may cause imbalance in oxidative stress levels and increased apoptosis. CGA improved Dex‐induced oxidative damage and apoptosis by reducing the activity of the ACE‐AngII‐AT1R pathway.

The animal experiment is divided into the exploratory and experimental phases, both of which use inbred SPF‐grade female 6‐week‐old C57BL/6 mice. The aforementioned experimental animals were purchased from Zhaoyan (Suzhou) New Drug Research Center Co., Ltd., and housed in the SPF‐grade experimental animal facility of Soochow University (SYXK(Su)2021‐0065). During the acclimation phase before the experiment, the mice had free access to water and food. This experiment meets the requirements and conditions for SPF‐grade animal housing and management, and all experimental animals and procedures have been approved by the Animal Ethics Committee of Soochow University and comply with national as well as international guidelines for the care and use of animals. CUS is a widely used experimental model in preclinical studies to simulate the effects of chronic stress on physiological and psychological outcomes. In this paradigm, rodents are exposed to a series of randomized, sporadic, and varied stressors selected from a pre‐determined set of interventions. This model is recognized for its ability to induce significant behavioral and physiological changes, such as HPA axis dysregulation, oxidative stress, and behavioral phenotypes associated with anxiety or depression [ 36 ]. In recent years, researchers have used CUS model animals to observe the damage of CUS to other organ functions [ 37 ]. In our study, the following 10 stressors were set, including food deprivation (no food for 12 h), water deprivation (no water for 12 h), forced swim (mice were placed in cages for 5 min with water at 25°C just enough for the mice to stand and not submerge their heads), cage tilting ( 45° cage‐tilt along the vertical axis for 6 h), social crowding (the same group of mice were placed in only one cage for 12 h), social isolation (a single mouse was housed per cage for 12 h), wet bedding (moistened sawdust for 6 h), restraint stress (mice were placed in a clean 50 mL conical tube with pierced holes for ventilation for 4 h), pinching tail (1 cm from the tip of the tail for 5 min by tweezer), and empty cage (absence of sawdust in cage for 6 h). Exploratory phase: The mice were randomly divided into the control group (no special treatment, Group C), low‐frequency stimulation group (CUS intervention every other day, Group L), medium‐frequency stimulation group (CUS intervention once daily, Group M), and high‐frequency stimulation group (CUS intervention twice daily, Group H). Experimental phase: The mice were randomly divided into the control group (no special treatment, Group C), high‐frequency stimulation group (CUS intervention twice daily, Group H), and high‐frequency stimulation + CGA group (CUS intervention twice daily with 100 mg/kg CGA (Aladdin, China) administered orally at the beginning of the fifth week after CUS intervention, Group A) [ 38 ], and the time intervention model diagrams in two phases were shown in Figures S1 and S2 . After a 7‐day acclimation period, the mice were subjected to the corresponding interventions according to their respective groups. The stressors had been mentioned above. Low, medium and high frequency CUS intervention lasted for 8 weeks. To prevent the mice from adapting to the stressors, the types of stressors for consecutive CUS interventions were different. However, during restraint interventions, control group mice were also deprived of food and water to eliminate confounding factors. Mice were weighed and recorded at a fixed time each week. After the interventions, blood and ovarian tissue samples were collected for subsequent experiments. The intervention schedules for both phases were administered randomly (see Tables S1 and S2 for detailed information). Dexamethasone (Dex, Pushitang, China) was dissolved in DMSO (Sigma, USA) and diluted to 1 µM with culture medium for cell experiments [ 3 ]. AngII (AbMole, USA) was dissolved in PBS and diluted to 10 µM with culture medium for cell experiments [ 27 ]. CGA (Aladdin, China) was dissolved in PBS and diluted to 10 µM with culture medium for cell experiments [ 39 ]. CS (Yuanye, China) was dissolved in PBS and diluted to 10 µM with culture medium for cell experiments [ 40 ]. Mouse blood samples were left standing for 2 h at room temperature, centrifuged at 2000 rpm for 15 min at 4°C to obtain serum samples. Obtained mouse serum and according to the manufacturer's instructions, ELISA kits (Jiangsu Meimian Industrial Co., Ltd, China) were used to detect serum levels of corticosterone (CORT, CORT ELISA Kit, MM‐0061M1), E 2 (E 2 ELISA Kit, MM‐0566M1), FSH (FSH ELISA Kit, MM‐45654M1), AMH (AMH ELISA Kit, MM‐44204M1), AngII (AngII ELISA Kit, MM‐0207M2), and Ang(1‐7) (Ang(1‐7) ELISA Kit, MM‐0754M2). After an acclimation training, the mice were fasted and deprived of water for 12 h after which the SCT was conducted. Each cage was provided with one bottle of drinking water and one bottle of 1% sucrose solution. Mice were allowed to drink freely, and after 24 h, the two bottles were removed (with bottle positions swapped every 6 h during the experiment to prevent positional bias), and the 24‐h sucrose preference percentage was calculated by an experimenter who was unaware of the experimental details. Sucrose preference percentage = (amount of 1% sucrose solution consumed / (amount of 1% sucrose solution consumed + amount of drinking water consumed)) × 100% [ 41 ]. Tape was applied to the last third of the mouse's tail and secured about 20 cm above the ground to keep the mouse inverted for 6 min, and the mouse's behavior during this period was recorded with a camera. Mice were given a 2‐min acclimation period before the experiment, and an experimenter who was unaware of the experimental details recorded the immobility time (when the mouse stopped struggling and remained still in the inverted position) from 3 to 6 min. The percentage of immobility time relative to the total experimental time was calculated [ 39 ]. The vaginal opening of mice was fully exposed. A sterile swab dipped in 0.9% NaCl solution was inserted into the vagina and rotated for one complete turn. The swab was then used to make a single‐layer smear on a glass slide, obtaining a vaginal exfoliative cell smear. After air‐drying the smear at room temperature, anhydrous methanol was added to fix the sample. The smear was then stained with hematoxylin and eosin (H&E, Yuanye, China), air‐dried again, and mounted with neutral resin. The exfoliative cell morphology was observed and recorded under an optical microscope (Nikon, Japan), and the number of days in each phase of the estrous cycle was documented. Ovarian tissue was fixed in 4% paraformaldehyde for 24 h, dehydrated, and cleared. The paraffin‐embedded ovarian tissue was sectioned into 8 µm histological slices, stained with H&E, and then mounted with neutral resin. The tissue morphology was observed under the microscope (Nikon, Japan). Follicles were counted by scanning each slice, with only those having a clear oocyte nucleus in the ovary being counted to avoid duplication [ 42 ]. Primordial follicles (oocytes surrounded by a single layer of flat GCs), primary follicles (oocytes surrounded by a single layer of cuboidal GCs), secondary follicles (oocytes surrounded by ≥ 2 layers of cuboidal GCs), mature follicles (follicular fluid increases and merges to form the follicular cavity), atretic follicles (oocytes degenerate and surrounded by multiple layers of dense GCs), and corpus luteum (collapsed follicle wall composed of GCs, connective tissue, and capillaries) [ 43 ]. Mouse ovarian tissues were isolated, homogenized in RIPA lysis buffer (Elabscience, China). After adjusting the protein concentration to be uniform, local ovarian AngII (AngII ELISA Kit, MM‐0207M2) and Ang(1‐7) (Ang(1‐7) ELISA Kit, MM‐0754M2) concentrations were measured according to the manufacturer's instructions using ELISA kits (Jiangsu Meimian Industrial Co., Ltd, China). Additionally, local ovarian GSH, CAT activity, and MDA levels were measured using assay kits (Nanjing Jiancheng, China) for GSH, CAT, and MDA. Ovarian tissue slices were obtained and stained according to the protocol of the apoptosis staining kit (Roche, USA). Image‐Pro Plus 6.0 (Media Cybernetics, Inc., USA) was used to select green fluorescent nuclei as the standard for positive cells in all images, and DAPI‐stained blue nuclei were used to identify total cells. Each image was analyzed to determine the number of positive cells and total cells, and the percentage of positive cells (positive cell count/total cell count × 100%) was calculated as the apoptosis rate (%). Apoptosis in KGN cells was assessed using an Annexin V‐FITC/PI apoptosis detection Kit (Beyotime, China) following the manufacturer's protocol. Briefly, KGN cells from each group were harvested, and the culture medium was aspirated. The cells were washed twice with pre‐cooled PBS, and 195 µL of Annexin V‐FITC binding buffer was added to each sample. Subsequently, 5 µL of Annexin V‐FITC and 10 µL of PI staining solution were added, and the mixture was gently vortexed to ensure even staining. The samples were incubated in the dark at room temperature for 15 min before analysis. Following 8‐week CUS intervention, mice were injected with 5 IU of pregnant mare serum gonadotropin (PMSG) (Aibei, China). After 46–48 h of stimulation, 5 IU of human chorionic gonadotropin (HCG) (Aibei, China) was administered intraperitoneally, and 14–16 h later, oocyte corona cumulus complexes (COCs) were collected from the oviducts. The COCs were moved to HEPES‐buffered M2 solution (Aibei, China), and then treated with 0.1% hyaluronidase (Aibei, China) for 1 min to remove the cumulus cells. The MII oocytes were washed thoroughly and transferred to M2 droplets under mineral oil (Aibei, China) at 37°C. COCs were obtained (using the same method as above), and 3–5 µL of sperm from 8‐week‐old male C57BL/6 mice, capacitated in TYH capacitation medium (Aibei, China) for 1 h, were combined with the COCs. The mixture was placed in HTF fertilization medium (Aibei, China) and incubated for 6 h in a cell culture incubator. Embryos were then transferred to KSOM embryo culture medium (Aibei, China) and cultured overnight in the incubator. The following day, the fertilization status of the oocytes was observed under the microscope. The KGN cell line, a human ovarian GC tumor cell line [ 44 ], was generously provided by Professor Li Chen (Chinese PLA General Hospital, NanJing, China). Cells were cultured using the method previously described [ 45 ], in Dulbecco's Modified Eagle's Medium (DMEM)/F12 medium (Gibco, USA) supplemented with 10% fetal bovine serum (Sigma, USA) and 0.5% penicillin‐streptomycin (Gibco, USA) at 37°C with 5% CO 2 . ROS levels in oocytes and KGN cells were measured using a ROS detection kit (Beyotime, China). MII oocytes were collected as described earlier. The DCFH‐DA probe was diluted to a final concentration of 10 µM in M2 solution, and the solution was incubated at 37°C with 5% CO 2 for 20 min. KGN cells were collected after applying the intervention drugs, and the DCFH‐DA probe was diluted to a final concentration of 10 µM in serum‐free cell culture medium and incubated at 37°C with 5% CO 2 for 20 min. Mitochondrial membrane potential levels in oocytes and KGN cells were measured using a mitochondrial membrane potential detection kit (Beyotime, China). MII oocytes were collected as described earlier. Incubated at 37°C with 5% CO 2 for 20 min after adding the working solution. KGN cells were collected after applying the intervention drugs, and the working solution was added to each well, followed by incubation at 37°C with 5% CO 2 for 20 min. CCCP in the kit was used as a positive control. Total RNA was extracted from ovarian tissues and cells using the RNA Quick Extraction Kit (Yishan, China) and quantified with the Qubit RNA HS Assay Kit (Invitrogen, CA). cDNA was synthesized using ABScript III Reverse Transcriptase (Abclonal, China) according to the manufacturer's instructions. qRT‐PCR was performed on the StepOnePlus system (Thermo Fisher Scientific, USA) using 2X Universal SYBR Green Fast qPCR Mix (Abclonal, China), and relative gene expression was calculated using the 2 −ΔΔCT method with GAPDH as the internal control. Primers, which were tested for specificity and efficiency, are listed in Table S3 . Proteins were extracted from ovarian tissues and cells using RIPA lysis buffer, and protein concentrations were measured using the BCA protein assay kit (Beyotime, China). Proteins were separated on SDS‐PAGE gels (Epizyme, China), transferred to PVDF membranes (Millipore, USA), and then blocked with 5% non‐fat milk at room temperature for 2 h. The following primary antibodies were used: AGT (Affinity, China, DF7976, 1:1000), AGTR1 (BOSTER, China, BM4949, 1:1000), AGTR2 (BOSTER, China, BM4557, 1:1000), MasR (Affinity, China, DF2818, 1:1000), ACE (Affinity, China, AF5197, 1:1000), ACE2 (Affinity, China, AF5165, 1:1000), BAX (Bimake, USA, A5131, 1:1000), Bcl‐2 (Affinity, China, BF9103, 1:1000), SOD2 (Selleck, USA, A5377, 1:1000), GPX4 (Selleck, USA, A5569, 1:1000), GAPDH (Proteintech, China, 60004‐1‐Ig, 1:10000). After overnight incubation at 4°C, the membranes were incubated with HRP‐conjugated secondary antibodies at room temperature for 1 h. Bands were detected using the ECL method, and the grayscale values were calculated using Image J software (NIH, USA), with GAPDH grayscale values used for normalization. Data are expressed as means ± standard deviations. Statistical analysis was performed using GraphPad Prism 8 (GraphPad Software, USA). Student's t test was used for comparing two groups and one‐way ANOVA for comparing three or more groups. A p value <0.05 was considered statistically significant. All experiments were independently repeated at least three times.

The OVRAS ACE‐AngII‐AT1R pathway may contribute to the decline of ovarian reserve function caused by chronic stress, CGA could help improve ovarian reserve function by reducing oxidative stress and apoptosis in ovarian tissue through downregulation of the ACE‐AngII‐AT1R pathway.

Our study reveals that CUS induces a decline in ovarian reserve function through the ACE‐AngII‐AT1R pathway within the OVRAS. CGA treatment effectively mitigates oxidative stress and apoptosis by downregulating this pathway, thereby alleviating the detrimental effects of chronic stress on ovarian reserve function. Ovarian aging is a global issue, and current clinical treatments and basic research are insufficient to meet real‐world needs, making it a research hotspot in reproductive medicine. The decline in ovarian reserve function, as a “predictive” marker for ovarian aging, has opened a new chapter in reproductive endocrinology research [ 54 ]. Accurately assessing ovarian reserve function and predicting the timing of ovarian aging is particularly important for career women who choose to delay marriage and childbirth. Long‐term stress from various sources, including increased social competition, excessive work load, interpersonal conflicts, and negative emotional stimuli, significantly impacts ovarian function and accelerates the decline in ovarian reserve function. The results of this study indicate that long‐term chronic stress leads to reduced AMH levels in mice, causes endocrine dysfunction in the ovaries, disrupted estrous cycles, and decreases the number of functional follicle, which is consistent with previous research findings [ 55 , 56 , 57 ]. Instead of directly inducing stress models via intravenous glucocorticoid injection, this study used the CUS intervention model for stress testing in mice. This model better simulates the unpredictable stressors encountered by reproductive‐age women in real life, providing a more accurate measurement of stress hormone levels under natural stress conditions. This method is superior to other approaches such as oral or injected CORT. Additionally, during the exploratory phase of this experiment, interventions at different frequencies (low, medium, and high) were used to reveal the impact of stress frequency on the decline in ovarian reserve function under chronic stress. Other studies have shown a close relationship between social psychological factors and infertility. There is an interactive cycle of promotion and constraint between the two [ 58 , 59 ]. Reduced ovarian reserve, one of the causes of infertility, often leads patients to choose assisted reproductive technologies (ART). However, reduced ovarian reserve can result in poorer pregnancy outcomes, which in turn increases the psychological burden on infertile women during treatment [ 60 , 61 ]. If not properly addressed, this can further lead to a vicious cycle of negative emotions and abnormalities in the HPA and HPO. Therefore, within the current biopsychosocial medical model, it is essential and urgent for healthcare providers to monitor both the physiological status and psychological well‐being of patients with reduced ovarian reserve. Like the classical systemic RAAS, the OVRAS includes two key peptides along with their enzymes and receptors that constitute two core pathways: ACE‐AngII‐AT1/2 R and ACE2‐Ang(1‐7)‐MasR. These pathways balance and regulate each other to maintain the relative stability of the ovarian microenvironment. OVRAS is involved in various important physiological functions such as folliculogenesis, regulation of ovulation, and steroid hormone synthesis [ 62 ]. Studies have shown that disruptions in OVRAS can lead to various ovarian diseases, including PCOS, ovarian hyperstimulation syndrome, and ovarian cancer [ 16 ]. Recent research by Jihyun Kim has shown that under chronic stress with high housing density, AngII and AGT expression in the OVRAS of mice are elevated, leading to a reduction in ovarian reserve function. However, no significant changes in ACE/AT1R expression were observed [ 27 ]. In this experiment, we used the CUS model to intervene in mice and observed that the mice exhibited certain anxiety and depressive tendencies in the SCT and the TST, while also assessing the expression of key components in the mouse OVRAS. We observed that under high‐frequency stimulation, the expression levels of Ace , Agt , and At1r in the OVRAS of mice were elevated at both the mRNA and protein levels. The ovarian tissue homogenate indicated an increase in AngII expression in the ovaries under medium and high‐frequency stimulation compared to the control group, suggesting excessive activation of the ACE‐AngII‐AT1R pathway in the OVRAS of mice subjected to chronic stress. Additionally, it is noteworthy that during in vivo experiments with low‐frequency stimulation, the expression levels of Ace2 , Ang(1‐7) and At2r were significantly increased compared to the control group, whereas no significant changes in the ACE2‐Ang(1‐7)‐MasR pathway were observed in KGN cells. We hypothesize that this phenomenon might be a result of negative feedback regulation in the ovaries in response to mild stress, which prevents excessive activation of the ACE‐AngII‐AT1R pathway and helps to maintain the balance of the ovarian microenvironment [ 63 , 64 ]. The follicle is the structural and functional unit of the ovary, consisting of the oocyte and GCs. GCs synthesize various hormones and growth factors to regulate the growth, differentiation, and maturation of oocytes, thereby controlling follicle development. Therefore, GCs are crucial for follicle activation and development, and their dysfunction is a key factor in follicular atresia. Abnormal oxidative stress is one of the significant causes of GC dysfunction [ 24 ]. Research indicates that elevated oxidative stress levels in GCs can contribute to several female reproductive disorders, including PCOS, endometriosis, and POF. Thus, decreasing oxidative stress in GCs may enhance oocyte quality and function. Oxidative stress is mainly caused by an imbalance between excessive ROS production and inadequate antioxidant defenses, which is regarded as a major factor leading to GC apoptosis. Prior research indicates that the extent of oxidative stress in oocytes largely depends on the antioxidant mechanisms provided by GCs, as the production levels of ROS are closely associated with oocyte‐cumulus cell interactions and the regulation of gamete function and development [ 65 ]. This study observed that chronic stress leads to abnormal elevation of ACE‐AngII‐AT1R axis activity in the OVRAS, which may be associated with increased oxidative stress levels in the ovaries. Additionally, increased ROS levels, reduced mitochondrial membrane potential, and decreased expression of ovarian antioxidant enzymes Sod2 and Gpx4 were observed in mouse oocytes. Dex‐induced oxidative stress and apoptosis levels increased in KGN cells, suggesting that an imbalance in ovarian oxidative stress levels may be related to the OVRAS [ 16 ]. Elevated MDA levels are associated with reduced activity of the antioxidant enzymes CAT and GSH. Oxidative stress arises from an imbalance between oxidative free radicals and antioxidants, with GSH and CAT playing crucial roles in the clearance of ROS. In this study, we also observed reduced antioxidant enzyme activity and increased MDA levels in the ovaries of mice under chronic stress, which is consistent with other studies. This suggests that chronic stress impairs ovarian function by increasing free radicals and decreasing antioxidant enzyme activity. Research has also indicated that oxidative stress can compromise mitochondrial structural integrity, resulting in cell apoptosis. Tunel apoptosis staining analysis revealed that the CUS group had more Tunel positive cells compared to the control group, with most of these cells being GCs surrounding the oocytes. Follicular growth and development are mainly regulated by GCs. Over‐apoptosis of GCs can speed up follicular depletion, manifesting as follicular atresia, which leads to DOR [ 48 ]. The Bcl‐2 protein prevents the release of pro‐apoptotic factors from mitochondria, thereby inhibiting apoptosis. BAX is a pro‐apoptotic protein crucial for mitochondrial membrane permeability and subsequent release of apoptotic molecules. The Bax/Bcl‐2 ratio may be more critical for determining apoptosis than the individual use of either protein. In this study, the Bax/Bcl‐2 ratio was increased in the chronic stress group of mice. Further addition of AngII to KGN cells elevated the Bax/Bcl‐2 ratio, suggesting that chronic stress may impair GC function and accelerate apoptosis via the ACE‐AngII‐AT1R pathway, thereby affecting oocyte function. CGA, a natural polyphenol found in coffee, fruits, and vegetables, exhibits established bioavailability in humans. However, its absorption is moderate due to complex metabolism primarily in the gastrointestinal tract and liver. It is absorbed via two mechanisms: rapid uptake in the stomach or upper gastrointestinal tract and slower absorption in the small intestine [ 29 ]. Toxicological, pharmacokinetic, and clinical studies in animals and humans have shown that oral CGA administration poses no significant toxicity or adverse effects, making it a safe dietary supplement within recommended dosages for healthy individuals. However, like other bioactive compounds, CGA may cause side effects at high doses or in susceptible populations. Rarely reported effects include gastrointestinal discomfort, mild laxative activity, and potential interactions with certain medications due to its influence on metabolic pathways [ 66 , 67 ]. The ovarian protective effects of CGA are primarily attributed to its anti‐inflammatory, antioxidant, and anti‐apoptotic properties, which collectively contribute to maintaining ovarian function and cellular integrity. Neda Abedpour discovered that culturing ovarian tissue in medium with 100 µM CGA could enhance follicle growth and development by lowering ROS levels in vitro [ 68 ]. Moreover, Mohd Zahoor ul Haq Shah reported that CGA could relieve letrozole‐induced PCOS symptoms in mice by elevating antioxidant enzyme expression levels in the ovaries [ 69 ]. Our research is the first to demonstrate that CGA improves ovarian reserve function by modulating the ACE‐AngII‐AT1R pathway in GCs. After chronic stress, mice exhibited excessive activation of the ACE‐AngII‐AT1R pathway in the OVRAS, leading to increased ovarian oxidative stress levels. After supplementing with CGA, the expression of ACE and AT1R decreased in the ovaries and GCs. ROS levels in oocytes and GCs were reduced, and mitochondrial membrane potential levels were restored. CGA contains ortho‐hydroxyl groups on its aromatic ring, which can scavenge ROS and alleviate lipid peroxidation [ 70 ]. In this study, the reduction in ovarian MDA levels following CGA treatment confirms previous findings. Additionally, antioxidant‐related indicators in the ovaries, such as GPX4, SOD2, CAT, and GSH, showed a certain degree of recovery in expression levels after CGA administration. These results indicate that CGA treatment reduced oxidative stress and apoptosis levels in the ovaries, alleviating ovarian damage induced by chronic stress. Based on existing evidence, we hypothesize that CGA downregulates the ACE‐AngII‐AT1R pathway through both direct and indirect mechanisms. Overactivation of this pathway elevates ROS production, disrupting ovarian oxidative stress balance, damaging GCs, and impairing the ovarian microenvironment. CGA enhances antioxidant activities, including SOD2 and GPX4, thereby reducing oxidative stress and limiting ROS production, which subsequently mitigates pathway activation. Additionally, CGA may modulate upstream oxidative stress signaling, downregulating ACE expression, reducing AngII production, and limiting AT1R binding, thereby suppressing downstream harmful effects. Emerging studies also suggest that polyphenols like CGA may regulate gene expression via epigenetic modifications [ 71 ]. CGA may epigenetically modulate genes linked to the ACE‐AngII‐AT1R pathway, thereby attenuating its activity in stress‐affected ovarian tissues. While additional studies are required to validate these mechanisms, our findings highlight the therapeutic potential of CGA in targeting this pathway. Chronic stress has been identified as a risk factor for DOR, with oxidative stress and apoptosis in GCs proposed as key contributors. However, the precise mechanisms linking chronic stress to impaired ovarian reserve remain poorly understood. Additionally, while antioxidant therapies have been investigated in other models of ovarian dysfunction, their efficacy in mitigating chronic stress‐induced DOR has been inadequately explored. To address these knowledge gaps, the present study focuses on two critical objectives: (1) elucidating the involvement of the OVRAS, particularly the ACE‐AngII‐AT1R axis, in the pathogenesis of chronic stress‐induced DOR and (2) evaluating the therapeutic potential of CGA, a dietary antioxidant, in mitigating oxidative stress and apoptosis associated with chronic stress. Our study underscores the significant impact of chronic stress on ovarian reserve function, primarily mediated through the ACE‐AngII‐AT1R pathway. These findings highlight the necessity of incorporating psychological evaluations into the clinical assessment of women with DOR. Addressing chronic stress as a modifiable risk factor should be a key component of a comprehensive treatment approach. Additionally, our results demonstrate the potential of CGA, an antioxidant, as a therapeutic adjunct to improve ovarian function by reducing oxidative stress and apoptosis in ovarian tissue. These findings suggest that CGA holds promise as a potential therapeutic agent for addressing DOR and infertility. Its ability to mitigate oxidative stress and apoptosis at the cellular level may enhance outcomes in ART, such as improving oocyte retrieval rates and embryo quality. Furthermore, CGA offers a non‐invasive, diet‐based strategy to complement existing treatments for women experiencing chronic stress or early signs of ovarian aging. We advocate for a multidisciplinary treatment strategy that integrates psychological support to alleviate chronic stress with targeted antioxidant interventions, such as CGA. This approach may offer a promising avenue to preserve ovarian reserve and improve reproductive health outcomes in women affected by stress‐induced DOR.

KGN cells, as an immortalized cell line, may not fully replicate the physiological conditions of GCs in vivo. This limitation could influence the accuracy of certain biological responses, particularly regarding cellular functions, hormone synthesis, and interactions with other cell types within the ovarian microenvironment. In addition, the variability in individual responses to CUS and CGA, influenced by genetic, environmental, and physiological factors, may lead to differences in the severity of DOR among mice. This inter‐individual variability could impact the generalizability of our findings. What's more, while our study demonstrated the protective effects of CGA on ovarian reserve function, we did not compare its efficacy with other antioxidants or therapeutic agents to highlight its relative advantages. Future studies should compare CGA with other antioxidants and compounds that alleviate oxidative stress and apoptosis to delineate its unique properties and optimize therapeutic strategies. Clinical trials assessing the safety, efficacy, and optimal dosage of CGA in women with DOR will be essential for its clinical translation. Additionally, further investigations into pathways implicated in ovarian dysfunction under chronic stress, including inflammation, autophagy, and mitochondrial dynamics, could uncover synergistic targets for intervention.

Ovarian reserve reflects the ability of follicles to develop and form oocytes and indirectly indicates ovarian function [ 1 , 2 ]. Diminished ovarian reserve (DOR) refers to the phenomenon where various physiological or pathological factors reduce the number and/or quality of oocytes in the ovary, resulting in decreased ovarian function. The characteristics of DOR include fewer antral follicles, elevated serum follicle‐stimulating hormone (FSH) levels, decreased serum estradiol (E 2 ), and anti‐müllerian hormone (AMH) concentrations [ 2 , 3 ]. DOR plays a vital role in the etiology of female infertility [ 4 ]. If left untreated, DOR may lead to premature ovarian insufficiency (POI) or even premature ovarian failure (POF) within a few years [ 5 , 6 ]. These conditions significantly impact for both reproductive and systemic health. Hormonal imbalances contribute to a spectrum of health issues, such as osteoporosis, cardiovascular diseases, and metabolic disorders. Moreover, the psychosocial burden is profound, including emotional distress, anxiety, and depression driven by infertility and the challenges of managing chronic health conditions. Therefore, studying the causes of DOR and implementing early interventions are crucial for preserving ovarian function and ensuring quality of life in women. Since Hans Selye's groundbreaking introduction of the “stress” theory in 1936, the significance of chronic stress in the onset and progression of diseases has been recognized [ 7 , 8 ]. Chronic stress, as one of the causes of DOR, has become increasingly important in modern society [ 3 ]. With the advancement of modern society, the pace of life has accelerated, and women of childbearing age are experiencing increasing pressures from work, study, and family. Prolonged and repeated chronic stress causes HPA dysfunction, a central neuroendocrine system that regulates the body's stress response and homeostasis, leading to abnormal secretion of glucocorticoids and body homeostasis [ 3 , 9 , 10 ]. Additionally, excessive activation of the HPA axis may impact the hypothalamic pituitary ovarian (HPO) axis [ 11 ], crucial for regulating female reproduction by controlling hormone secretion for ovarian follicle development, ovulation, and menstrual cycles. Elevated serum cortisol levels from various stressors disrupt gonadotropin release and steroid hormone biosynthesis, thus impacting follicle maturation and ovulation in the ovary [ 12 , 13 ]. Nevertheless, studies on the adverse impacts and underlying mechanisms of increased glucocorticoids from chronic stress on ovarian function and follicle development remain limited. The classic renin‐angiotensin‐system (RAS) is a key humoral regulatory system, traditionally linked to the development and progression of cardiovascular, renal, and endocrine diseases [ 14 ]. Recent research indicates that beyond the classic endocrine RAS, RAS is also expressed in many organs, including the ovaries [ 15 ]. The OVRAS is an intrinsic hormonal signaling network within the ovary, essential for regulating follicular development, ovulation, and maintaining ovarian vascular integrity and cellular homeostasis. It primarily consists of two key signaling pathways: ACE‐AngII‐AT1R and ACE2‐Ang(1‐7)‐MasR, which regulate ovarian function, folliculogenesis, development, atresia, and the development of ovarian diseases [ 16 , 17 , 18 , 19 ]. Research has demonstrated that an imbalance in the two signaling pathways is linked to folliculogenesis and ovulation disorders, possibly involving a local oxidative stress imbalance in the ovaries and increased apoptosis [ 20 , 21 ]. Oxidative stress imbalance is recognized as one of the pathogenic factors associated with DOR [ 22 ]. Numerous studies indicate that oxidative stress contributes to impairing ovarian function, poor oocyte quality, and ovulation disorders in contemporary women [ 23 , 24 ], as excessive reactive oxygen species (ROS) can disrupt the balance of the body's oxidative and antioxidant systems. Research has shown that chronic stress elevates oxidative stress levels in the body. In the female reproductive system, oxidative stress impacts several stages, including primordial follicle formation, follicle development, maturation, and atresia, and contributes to the development of other ovarian disease etiologies, such as apoptosis of GCs, mitochondrial dysfunction, and calcium ion imbalance [ 25 , 26 , 27 ]. Thus, lowering oxidative stress levels might be a crucial strategy for enhancing follicle reserve and developmental capacity in DOR patients. Chlorogenic acid (CGA), formed by the esterification of caffeic acid and quinic acid, is one of the most abundant natural polyphenols found in coffee, fruits, and vegetables [ 28 ]. Previous studies have shown that CGA possesses various biological functions, including antiangiogenesis, anticancer, and antiglycation properties under metabolic disorders [ 29 , 30 ]. Additionally, CGA plays a key role in oxidative stress reduction by scavenging ROS, alleviating oxidative damage, and mitigating mitochondrial dysfunction through various pathways. Studies have found CGA could improve duodenal ulcer injury caused by chronic stress by inhibiting IL‐6/JAK2/STAT3 signaling pathway [ 31 ]. Meanwhile, we found that chlorogenic acid could regulate the RAS by inhibiting the activity of ACE in plasma or tissues, thereby reducing blood pressure [ 32 , 33 ]. Other studies have shown that CGA could improve follicle development by reducing oxidative stress levels in the ovaries of polycystic ovary syndrome (PCOS) rats [ 34 ] and could reduce apoptosis levels in GCs, thereby treating ovarian granulosa cell (GC) damage induced by zearalenone in mice [ 35 ]. However, whether CGA could improve DOR caused by chronic stress by regulating OVRAS and the mechanisms involved remains limited. In this study, we aim to elucidate the role of the OVRAS in DOR and investigate antioxidant stress and anti‐apoptotic effects of CGA in chronic unpredictable stress (CUS)‐induced DOR, and its underlying molecular mechanisms.

The authors declare no conflicts of interest.

Supporting Information