Does diminished ovarian reserve impact oxidation–reduction potential in follicular fluid?

Abstract

Objective

This study aimed to evaluate oxidative stress in follicular fluid by measuring oxidation–reduction potential and determine its correlation with embryo quality in patients with diminished ovarian reserve.

Methods

This cross-sectional study included 134 patients with diminished ovarian reserve and 82 patients with normal ovarian reserve. Follicular fluid samples were collected from each patient via transvaginal aspiration under ultrasound guidance. Oxidation–reduction potential was measured using the Male Infertility Oxidative system, following the manufacturer’s protocol.

Results

The levels of anti-Müllerian hormone, number of oocytes retrieved, rate of blastocyst formation, and rate of high-quality embryos were significantly lower in the diminished ovarian reserve group than in the normal ovarian reserve group. Correlation analysis revealed that oxidation–reduction potential values had a negative correlation with the number of oocytes retrieved (r = −0.262, p = 0.002), rate of mature oocytes (r = −0.260, p = 0.002), and rate of normal fertilization (r = −0.207, p = 0.016); however, these values showed a positive correlation with the duration of ovarian stimulation (r = 0.186, p = 0.031).

Conclusions

Oxidative stress in follicular fluid correlates with a small number of oocytes retrieved, lower proportion of mature oocytes, and lower rate of normal fertilization in patients with diminished ovarian reserve.

Keywords

Diminished ovarian reserve oxidation–reduction potential oxidative stress follicular fluid Male Infertility Oxidative system in vitro fertilization outcomes

Background

While using assisted reproductive techniques, high-quality oocytes are essential for improving fertilization rates and embryo outcomes, thus optimizing in vitro fertilization (IVF) outcomes.¹ Follicular fluid (FF) is an ideal source of noninvasive biochemical markers for oocyte development. FF is found in the follicular antrum and is pivotal in follicular development and oocyte competence. It is produced by the secretions of theca cells, granulosa cells, and oocytes, along with the diffusion of plasma components across the blood follicle barrier. FF contains a wide range of molecules, including microRNAs, polysaccharides, lipids, cytokines, inflammatory mediators, growth factors, antioxidant enzymes, and reactive oxygen species (ROS).^2–4

Oxidative stress (OS) arises from an imbalance between ROS and the antioxidant defense system, potentially leading to oxidative damage of DNA, proteins, and lipids. The impact of OS in FF on oocyte and embryo quality and IVF outcomes remains debatable. Agarwal et al. proposed that OS affects oocyte and embryo quality, impacting fertilization.⁵ However, Jozwik et al. suggested that OS markers in FF do not necessarily reflect the reproductive potential of oocytes.⁶ Other studies have shown that women who conceived through IVF had higher levels of ROS and lipid peroxidation than those who did not.⁷ Conversely, Oyawoye et al. indicated that higher total antioxidant capacity may improve fertilization potential in patients undergoing IVF treatment.⁸

Diminished ovarian reserve (DOR) is characterized by a reduction in ovarian reproductive capacity, which results from decreased oocyte quantity or quality, thereby impairing fertility. Factors contributing to DOR include age, chemoradiation, environmental factors, genetic predispositions, and immune system alterations.⁹ Previous studies have demonstrated that in several gynecological conditions, such as polycystic ovary syndrome (PCOS) and endometriosis, the quality of oocytes and embryos is closely associated with OS.^10–12 However, limited studies have focused on OS markers in the FF of patients with DOR.

The relationship between OS in the FF of DOR patients and embryo quality warrants further investigation. This study explored the association between OS, as measured by oxidation–reduction potential (ORP), and DOR.

Methods

Study design

This cross-sectional study was conducted at the Hue Center for Reproductive Endocrinology and Infertility, Hue University of Medicine and Pharmacy Hospital, Vietnam, from February 2023 to October 2024. The study was approved by the Ethics Committee of Hue University of Medicine and Pharmacy (approval number H2023/016, dated 20 February 2023). Participants were recruited consecutively from our center. Written informed consent was obtained from all participants, and all patient details were de-identified to ensure confidentiality. The study was conducted following the Declaration of Helsinki, as revised in 2024.

All patients included in this study were indicated for IVF. FF samples were obtained on the day of oocyte retrieval from individuals with DOR (DOR, n = 134) and those without ovarian disorders (normal ovarian reserve (NOR), n = 82). The exclusion criteria included oocyte or sperm donors.

This study was reported following the Strengthening the Reporting of Observational studies in Epidemiology (STROBE) guidelines.¹³

Ovarian stimulation and oocyte retrieval

Patients undergoing IVF or intracytoplasmic sperm injection (ICSI) cycles were indicated to undergo basal ultrasound on days 2–3 of their menstrual cycle to assess the antral follicle count and exclude the presence of functional ovarian cysts. Serum estradiol and follicle-stimulating hormone (FSH) levels were measured on days 2–3 of the menstrual cycle using an automated immunoassay analyzer. The predefined cutoff values for FSH and estradiol were 10 mIU/mL and 70 pg/mL, respectively. The diagnosis of DOR was established based on the Bologna criteria.

The participants underwent controlled ovarian stimulation using a gonadotropin-releasing hormone (GnRH) antagonist protocol and recombinant FSH (follitropin alfa). Subsequently, 225–300 IU of recombinant FSH (Gonal F®, Merck KGaA, Darmstadt, Germany) was administered from day 2 or 3 until the trigger day. GnRH antagonist (Cetrotide® 0.25 mg, Merck, Germany) was introduced once the follicles reached a diameter of 14 mm. Recombinant human chorionic gonadotropin (Ovitrelle®, 250 mcg, Merck, Germany) was administered when at least two preovulatory follicles with 16–22 mm diameter were observed. Oocyte retrieval was performed under transvaginal ultrasound guidance 36 h after the ovulation trigger.

Sperm DNA fragmentation assessment

Sperm DNA fragmentation was evaluated using the Halosperm R10 kit, following the manufacturer’s instructions. This technique relies on the principle that sperm with intact DNA form large- or medium-sized halos of dispersed chromatin after denaturation and nuclear protein removal, whereas fragmented DNA results in small halos, no halos, or degenerated sperm.

Semen samples were processed according to the kit protocol, which included dilution and embedding in agarose gel. The samples were then subjected to lysis for the removal of nuclear proteins, followed by dehydration and staining for analysis. An automated imaging system was used to assess DNA fragmentation by measuring the halo size to determine the DNA fragmentation index (DFI). Sperm with small halos, those with no halos, and those showing degeneration were classified as fragmented. A DFI >15% indicated sperm DNA fragmentation, which may negatively impact fertilization and embryo development. All analyses were conducted under standardized laboratory conditions to ensure accuracy and reproducibility.

Collection of FF and measurement of ORP

FF samples were carefully aspirated from the first follicles under vaginal ultrasound guidance. To ensure analytical accuracy, FF samples were processed immediately after aspiration to measure ORP, with strict precautions taken to avoid contamination from flushing media or blood. All samples were analyzed fresh, without being subjected to freezing.

ORP measurements were performed using the Male Infertility Oxidative system (MiOXSYS) (Caerus Biotechnologies, Geneva, Switzerland), following the manufacturer’s standardized protocol. This electrochemical system employs a silver/silver chloride (Ag/AgCl) reference electrode to quantify static ORP (sORP), expressed in millivolts. The sORP reflects the overall redox balance within the sample, representing the net interaction between total oxidants (ROS) and antioxidants present in the FF, thereby serving as a robust biomarker of OS.

Each FF sample was applied directly onto a sensor and analyzed in real-time using the MiOXSYS device, which automatically computes the ORP values without the need for manual calculations or external calibration. The analyzer was readily available in the oocyte retrieval suite, enabling immediate assessment without delays from storage or sample transport.

Sperm preparation and ICSI

Before ICSI, sperm were prepared using the Sil-select PlusTM density gradient system (Fertipro®, Beernem, Belgium) and rinsed twice with SpermRinse (Vitrolife®, Sweden). The final sperm suspension was resuspended in 0.2–0.5 mL of SpermRinse medium.

All patients underwent routine ICSI cycles. Cumulus–oocyte complexes were extracted and washed with G-MOSP PLUS (Vitrolife®, Sweden) and then cultured in G-IVF PLUS (Vitrolife®, Sweden) at 37°C for 2 h in an incubator under 6.0% CO₂ and 5.0% O₂. Subsequently, 80 IU of Hyase (Vitrolife®, Sweden) was used to induce denudation.

Post-ICSI, all mature injected oocytes were cultured individually in G-TL (Vitrolife®, Sweden) and covered with Ovoil (Vitrolife®, Sweden) at 37°C under 6.0% CO₂ and 5.0% O₂. Embryos were cultured for 5 days in a benchtop incubator. All embryos were evaluated according to the Istanbul consensus. Good- and medium-quality embryos were selected for cryopreservation using vitrification.

Statistical analysis

Data were analyzed using IBM SPSS Statistics for Windows Version 23.0 (IBM Corp., Armonk, NY, USA). Data were presented as mean ± standard deviation. Differences between the DOR and NOR groups were assessed using the Student’s t-test for continuous variables and the Mann–Whitney U test for categorical variables. Correlation analysis was performed to explore the relationship between ORP and embryo quality. A p-value <0.05 was considered to indicate statistical significance.

Results

In total, 216 samples were collected from 134 patients with DOR and 82 patients with NOR. The characteristics of the DOR and NOR groups are summarized in Table 1. DOR patients were significantly older than NOR patients (p = 0.004). Patients with DOR showed increased FSH levels (7.31 vs. 6.17 mIU/mL, p = 0.001), higher body mass index (20.76 vs. 19.44, p = 0.002), and greater total FSH dosage (2392.93 vs. 1752.88 IU, p < 0.001) than those with NOR. Additionally, anti-Müllerian hormone levels were significantly lower in the DOR group than in the NOR group (1.16 vs. 3.02 ng/mL, p < 0.001).

Table 1.

Baseline indicators of patients with DOR and those with NOR.

Factors	NOR (n = 82)	DOR (n = 134)	p-value
Female age (years)^&	34.70 ± 4.03	35.71 ± 5.36	0.004*
Male age (years)^&	37.26 ± 5.32	38.27 ± 6.47	0.055
BMI (kg/m²)^&	19.44 ± 5.74	20.76 ± 2.25	0.002*
AMH (ng/mL)^#	3.02 (2.04–7.57)	1.16 (0.00–6.48)	0.000*
AFC^&	14.06 ± 4.17	6.69 ± 4.34	0.511
FSH (mIU/mL)^#	6.17 (2.96–48.00)	7.31 (1.00–22.93)	0.001*
LH (mIU/mL)^#	5.70 (1.00–13.00)	5.10 (0.00–27.00)	0.137
E2 (pg/mL)^#	35.29 (12.34–79.11)	40.86 (12.58–280)	0.000*
Total FSH used (IU)^&	1752.88 ± 783.02	2392.93 ± 536.86	0.000*
Days of stimulation^# (days)	9.00 (6.00–12.00)	9.00 (6.00–13.00)	0.559
DFI (%)	17.18 (6.0–37.8)	17.00 (5.2–35.0)	0.810

DOR: diminished ovarian reserve; NOR: normal ovarian reserve; BMI: body mass index; AMH: anti-Müllerian hormone; AFC: antral follicle count; FSH: follicle-stimulating hormone; LH: luteinizing hormone; E2: estradiol; DFI: DNA fragmentation index.

*Difference is considered significant.

^&Values are presented as mean ± SD for continuous indicators following a normal distribution. Student’s t-test was used to compare continuous variables.

^#Values are presented as median (interquartile range) for continuous indicators following a non-normal distribution. Mann–Whitney U test was used to compare categorical variables.

There was no statistically significant difference in sperm DFI between the NOR and DOR groups. The mean DFI in the NOR group was 17.18% (range: 6.0%–37.8%), while that in the DOR group was 17.00% (range: 5.2%–35.0%) (p = 0.810).

Laboratory results are presented in Table 2. The ORP value in the DOR group was significantly higher than that in the NOR group (102.40 vs. 61.03, p = 0.044). The number of oocytes retrieved was lower in patients with DOR than in those with NOR (6.05 vs. 14.74, p < 0.001). The rates of mature oocytes and normal fertilization were comparable between the two groups (p = 0.114 and p = 0.875, respectively). Furthermore, the ORP value was significantly elevated in the DOR group (102.40 vs. 61.03, p = 0.044). The blastocyst formation rate and high-quality embryo rate were lower in the DOR group (53.13% vs. 68.74%, p = 0.001 and 32.54% vs. 47.04%, p = 0.014) than in the NOR group.

Table 2.

ORP values and embryo quality in the DOR and NOR groups.

Characteristics	NOR (n = 82)	DOR (n = 134)	p-value
ORP (mV)^&	61.03 ± 24.76	102.40 ± 21.37	0.044*
pH^&	7.57 ± 0.27	7.29 ± 1.14	0.070*
Number of oocytes retrieved^#	12.00 (7–46)	6.00 (1–19)	0.000*
Mature oocytes rate (%)^&	76.89 ± 17.50	87.34 ± 32.33	0.114
Normal fertilization rate (%)^#	76.39 (14.29–100)	75.00 (0–100)	0.773
Blastocyst formation rate (%)^#	80.00 (14.29–100)	62.50 (0–100)	0.001*
High-quality embryo rate (%)^&	47.04 ± 25.00	32.54 ± 31.08	0.014*

ORP: oxidation–reduction potential; DOR: diminished ovarian reserve; NOR: normal ovarian reserve.

Difference is considered significant.

Values are presented as mean ± SD for continuous indicators following a normal distribution. Student’s t-test was used to compare continuous variables.

Values are presented as median (interquartile range) for continuous indicators following a non-normal distribution. Mann–Whitney U test was used to compare categorical variables.

Table 3 shows the correlation between ORP values, clinical markers, and embryo quality in the DOR group. ORP levels were negatively correlated with the number of oocytes retrieved (r = −0.262, p = 0.002), rate of mature oocytes (r = −0.260, p = 0.002), and rate of normal fertilization (r = −0.207, p = 0.016). The duration of stimulation was positively correlated with ORP levels (r = 0.186, p = 0.031).

Table 3.

Correlation between ORP values and clinical indicators. Embryo quality in the DOR group.

Characteristics	r	p-value
Female age (years)	0.133	0.125
AFC	−0.156	0.072
Female BMI (kg/m²)	0.142	0.102
AMH (ng/mL)	−0.078	0.647
FSH (mIU/mL)	0.085	0.330
E2 (pg/mL)	0.092	0.290
LH (mIU/mL)	−0.064	0.461
Days of stimulation	0.186	0.031*
Total FSH dosage (IU)	0.217	0.012*
Number of oocytes retrieved	−0.262	0.002*
Mature oocytes rate (%)	−0.260	0.002*
Normal fertilization rate (%)	−0.207	0.016*
Blastocyst formation rate (%)	−0.100	0.250
High-quality embryo rate (%)	−0.095	0.275

DOR: diminished ovarian reserve; NOR: normal ovarian reserve; ORP: oxidation–reduction potential; AFC: antral follicle count; BMI: body mass index; AMH: anti-Müllerian hormone; FSH: follicle-stimulating hormone; E2: estradiol: LH: luteinizing hormone.

Discussion

This study revealed a significant negative correlation between OS levels in FF and several critical outcomes in patients with DOR, including the number of oocytes retrieved, proportion of mature oocytes, and rate of normal fertilization. These results underscore the importance of redox balance in FF as a determinant of oocyte developmental competence. Furthermore, while the relationship between OS and blastocyst formation or embryo quality was less definitive, the data suggest that elevated OS may preclude embryo development.

Our findings align with existing literature highlighting the detrimental effects of ROS on oocyte quality. Previous studies have shown that ROS adversely impact oocyte quality by accelerating apoptosis, compromising spindle integrity, inducing chromosomal aneuploidy, causing DNA damage, and disrupting cytoskeletal structure.¹⁴ Terao et al. suggested that the oxidation–reduction equilibrium in FF during oocyte retrieval is crucial for fertilization and embryonic division.¹⁵ Reduced OS might enhance fertilization potential and increase the rate of high-quality embryos. Several previous publications have also reported the negative impact of high ROS in FF on fertilization rate.^16,17 Moreover, the concentration of glutathione, a principal antioxidant, in FF was diminished in patients exhibiting low fertilization rates during IVF/ICSI cycles, along with a reduced rate of high-quality blastocysts.¹⁶ In women with PCOS, elevated total oxidant capacity could be associated with a reduced rate of high-quality embryos on day 3, subsequently leading to lower blastocyst formation rates (p < 0.05).¹¹ Our study adds to this body of evidence by focusing on DOR patients, a cohort that has been less explored in this context. Clinically, Fabjan et al. concluded that there is a substantial positive correlation between estradiol levels and total antioxidant capacity in FF (r = 0.26).¹⁷ Oxidation–reduction balance significantly influenced estradiol synthesis and steroid hormone production from granulosa cells.

The mechanism by which OS adversely affects oocyte quality remains unclear. OS might also negatively affect oocytes through the mechanism related to ovarian aging because follicular atresia is initiated by the loss of theca and granulosa cells,¹⁸ primarily due to chromosomal segregation errors during meiotic prophase.¹⁹ In our investigation, OS in aging ovarian FF was negatively associated with the number of oocytes retrieved, rate of mature oocytes, and successful fertilization. In addition to steroid hormones and granulosa cells, the FF contains cytokines, macrophages, and leukocytes, which produce ROS. The OS in the FF may either be attributed to the impaired metabolism of the oocyte or the environment. The OS could promote lipid peroxidation energy expenditure and affect the cell nucleus.²⁰ All of these phenomena might have adverse effects on oocyte maturation and fertilization capacity. However, the detrimental impact of OS on the rate of blastocyst formation and rate of high-quality embryos remains unclear. We hypothesize that oocytes capable of normal fertilization and embryogenesis may naturally overcome significant OS. When OS is elevated, oocyte DNA is compromised beyond the repair capacity at the pronuclear stage, and such oocytes may become incapable of fertilization.²¹

Measuring OS markers in FF could become a practical adjunctive method for assessing oocyte quality and tailoring stimulation protocols, particularly in patients with DOR. Antioxidant administration during IVF cycles may help improve oocyte quality, enhance IVF outcomes, and increase the success rate of treatment cycles. The mechanisms of action of antioxidants are known; they can enhance mitochondrial function or scavenge free radicals. Antioxidant supplementation could be prescribed as an oral pretreatment in patients undergoing IVF cycles.²² Further studies are needed to evaluate the positive effect of antioxidants on oocyte quality, IVF outcomes, and OS levels. However, there is no consensus on the treatment duration and the dosage of antioxidant supplementation. This emphasizes the need for more data for the clinical application of antioxidant protocols in patients undergoing IVF cycles. Antioxidants can be administered as an adjuvant therapy in the media during in vitro culture.²² According to Hosseini et al., supplementation with exogenous antioxidants such as βME can help increase the neutralization of embryos to ROS. This effect would be optimized if the supplementation is conducted throughout the culture period.²³ Adding ascorbic acid and rosmarinic acid as antioxidant substances in culture media could lead to a better expansion of cumulus cells, especially in class 1 and class 2 cumulus–oocyte complexes.²⁴ The use of antioxidants has several benefits in improving oocyte quality, increasing the success rate of IVF treatment cycles through oral administration or addition to the culture medium. Identifying the necessary target patient group for antioxidant indication is very important, especially in cases of reduced ovarian reserve.

A notable strength of this study is the use of a standardized method for measuring OS, allowing reliable quantification of the redox state in FF. Additionally, the MiOXSYS system provides rapid, precise, and minimally invasive assessment of OS, enhancing the efficiency of data collection. This is one of the first studies in Vietnam to evaluate OS in FF using the ORP method in patients with reduced ovarian reserve. However, some limitations of this study must be acknowledged. This study was conducted at a single center, which may limit the generalizability of the findings to the broader Vietnamese population. In addition to OS, IVF outcomes can be affected by various confounding factors, including female patient age, male partner characteristics, and laboratory embryo culture conditions. In our analysis, female patient age was included as a covariate in a multivariate regression model. However, its impact did not reach statistical significance. We acknowledge that the limited sample size, coupled with the presence of multiple confounders, may have reduced the statistical power to detect age-related effects on IVF outcomes in this cohort.

In terms of male factors, although the integrity of sperm DNA is known to be a critical factor in embryo development, our findings showed no significant difference in the sperm DFI between the NOR and DOR groups. This suggests that sperm DNA quality was relatively comparable between the two groups and thus was unlikely to be a confounding factor in the observed differences between oocyte and embryo outcomes. Given that all patients underwent ICSI, which allows for the selection of motility and morphologically normal sperm, the potential influence of DFI may have been further minimized. Therefore, the poorer embryo quality observed in the DOR group is more likely attributed to oocyte-related factors rather than sperm DNA integrity. These findings support the interpretation that DOR is associated with compromised oocyte quality, independent of sperm-related parameters.

An additional consideration is the timing of sperm DFI assessment. In our study, DFI testing was conducted during the initial infertility workup, prior to the initiation of treatment cycles. We did not perform DFI evaluation on the day of ICSI. Although same-day DFI analysis could potentially offer more relevant and dynamic insights into the gamete microenvironment, this approach was not pursued to streamline clinical operations and minimize the financial burden for patients, given the relatively high cost of DFI testing. We recognize this as a limitation of our study and intend to conduct DFI assessments on the day of ICSI in future research to enhance data precision and clinical applicability.

Conclusion

OS in FF correlates with fewer retrieved oocytes, reduced maturity rates, and decreased normal fertilization rates in patients with DOR. These findings suggest that the levels of ORP in FF can predict IVF outcomes in patients with DOR.

Supplemental Material

sj-pdf-1-imr-10.1177_03000605251349926 - Supplemental material for Does diminished ovarian reserve impact oxidation–reduction potential in follicular fluid?

Supplemental material, sj-pdf-1-imr-10.1177_03000605251349926 for Does diminished ovarian reserve impact oxidation–reduction potential in follicular fluid? by Minh Tam Le, Nguyen Dac Nguyen, Hong Nhan Thi Dang and Quoc Huy Vu Nguyen in Journal of International Medical Research

Footnotes

Acknowledgement

None.

Author contributions

MTL, NDN, and HNTD developed the study concept and designed the study; HNTD and NDN acquired the data for analysis; HNDT performed statistical analysis; MTL, NDN, and HNDT drafted the first version of the manuscript; all authors contributed to the interpretation of the data and provided critical revision for important intellectual content and approved the final manuscript.

Data availability statement

The data supporting this study’s findings are available on request from the corresponding author. The data are not publicly available due to privacy or ethical restrictions.

Declaration of conflicting interests

The authors declare no potential conflicts of interest.

Ethics approval and consent to participate

This study was approved by the Hue University of Medicine and Pharmacy Ethics Committee (approval number H2023/016). Written informed consent was obtained from the patients for the publication of this study.

Funding

This work was supported by the Hue University project funding (grant number DHH2023-04-198) and the Core Research Program of Hue University (grant number NCTB.DHH.2025.07; Research Group on Reproductive Medicine). The funders did not influence the content of the publication.

ORCID iD

Minh Tam Le

References

Wiener-Megnazi

Auslender

Dirnfeld

, et al. Oxidative stress indices in follicular fluid as measured by the thermochemiluminescence assay correlate with outcome parameters in in vitro fertilization. Fertil Steril 2004; 82: 1171–1176.

Freitas

Neto

Matos

, et al. Follicular fluid redox involvement for ovarian follicle growth. J Ovarian Res 2017; 10: 44.

Nejabati

Roshangar

Nouri

Follicular fluid extracellular vesicle miRNAs and ovarian aging. Clin Chim Acta 2023; 538: 29–35.

Niu

Xia

, et al. Follicular fluid cytokine composition and oocyte quality of polycystic ovary syndrome patients with metabolic syndrome undergoing in vitro fertilization. Cytokine 2017; 91: 180–186.

Agarwal

Gupta

Sharma

Oxidative stress and its implications in female infertility – a clinician’s perspective. Reprod Biomed Online 2005; 11: 641–650.

Jozwik

Wolczynski

Jozwik

, et al. Oxidative stress markers in preovulatory follicular fluid in humans. Mol Hum Reprod 1999; 5: 409–413.

Pasqualotto

Agarwal

Sharma

, et al. Effect of oxidative stress in follicular fluid on the outcome of assisted reproductive procedures. Fertil Steril 2004; 81: 973–976.

Oyawoye

Gadir

Garner

, et al. Antioxidants and reactive oxygen species in follicular fluid of women undergoing IVF: relationship to outcome. Hum Reprod 2003; 18: 2270–2274.

Park

Walsh

Berkowitz

KM.

Mechanisms of ovarian aging. Reproduction 2021; 162: R19–R33.

10.

Gongadashetti

Gupta

Dada

, et al. Follicular fluid oxidative stress biomarkers and ART outcomes in PCOS women undergoing in vitro fertilization: a cross-sectional study. Int J Reprod Biomed 2021; 19: 449–456.

11.

Liu

Zhao

, et al. Oxidative stress markers in the follicular fluid of patients with polycystic ovary syndrome correlate with a decrease in embryo quality. J Assist Reprod Genet 2021; 38: 471–477.

12.

Nasiri

Moini

Eftekhari-Yazdi

, et al. Oxidative stress statues in serum and follicular fluid of women with endometriosis. Cell J 2017; 18: 582–587.

13.

Von Elm

Altman

Egger

; STROBE Initiativeet al. The Strengthening the Reporting of Observational Studies in Epidemiology (STROBE) statement: guidelines for reporting observational studies. Ann Intern Med 2007; 147: 573–577.

14.

Lin

Wang

Oxidative stress in oocytes and embryo development: implications for in vitro systems. Antioxid Redox Signal 2020; 34: 1313–1331.

15.

Terao

Wada-Hiraike

Nagumo

, et al. Role of oxidative stress in follicular fluid on embryos of patients undergoing assisted reproductive technology treatment. J Obstet Gynaecol Res 2019; 45: 1884–1891.

16.

Sabatini

Wilson

Lower

, et al. Superoxide dismutase activity in human follicular fluid after controlled ovarian hyperstimulation in women undergoing in vitro fertilization. Fertil Steril 1999; 72: 1027–1034.

17.

Fabjan

Vrtačnik-Bokal

Kumer

, et al. Determination of oxidative stress balance in follicular fluid. J Lab Med 2018; 42: 51–58.

18.

Nishihara

Matsumoto

Hosoi

, et al. Evaluation of antioxidant status and oxidative stress markers in follicular fluid for human in vitro fertilization outcome. Reprod Med Biol 2018; 17: 481–486.

19.

Appasamy

Jauniaux

Serhal

, et al. Evaluation of the relationship between follicular fluid oxidative stress, ovarian hormones, and response to gonadotropin stimulation. Fertil Steril 2008; 89: 912–921.

20.

Alam

Oxidative stress and oocyte microenvironment. In: Subfertility. Elsevier, 2020, pp. 175–184.

21.

Matsuda

Inoue

Goto

, et al. cFLIP regulates death receptor-mediated apoptosis in an ovarian granulosa cell line by inhibiting procaspase-8 cleavage. J Reprod Dev 2008; 54: 314–320.

22.

Rodríguez-Varela

Labarta

Clinical application of antioxidants to improve human oocyte mitochondrial function: a review. Antioxidants (Basel) 2020; 9: 1197.

23.

Hosseini

Forouzanfar

Hajian

, et al. Antioxidant supplementation of culture medium during embryo development and/or after vitrification-warming; which is the most important? J Assist Reprod Genet 2009; 26: 355–364.

24.

Boldura

Marc

Otava

, et al. Utilization of rosmarinic and ascorbic acids for maturation culture media in order to increase sow oocyte quality prior to IVF. Molecules 2021; 26: 7215.

Supplementary Material

Please find the following supplemental material available below.

For Open Access articles published under a Creative Commons License, all supplemental material carries the same license as the article it is associated with.

For non-Open Access articles published, all supplemental material carries a non-exclusive license, and permission requests for re-use of supplemental material or any part of supplemental material shall be sent directly to the copyright owner as specified in the copyright notice associated with the article.

0.00 MB

0.12 MB